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PHARMACOLOGY AND PATHOPHYSIOLOGY OF THE CONTROL OF BREATHING
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LUNG BIOLOGY IN HEALTH AND DISEASE Executive Editor Claude Lenfant Former Director, National Heart, Lung, and Blood Institute National Institutes of Health Bethesda, Maryland
1. Immunologic and Infectious Reactions in the Lung, edited by C. H. Kirkpatrick and H. Y. Reynolds 2. The Biochemical Basis of Pulmonary Function, edited by R. G. Crystal 3. Bioengineering Aspects of the Lung, edited by J. B. West 4. Metabolic Functions of the Lung, edited by Y. S. Bakhle and J. R. Vane 5. Respiratory Defense Mechanisms (in two parts), edited by J. D. Brain, D. F. Proctor, and L. M. Reid 6. Development of the Lung, edited by W. A. Hodson 7. Lung Water and Solute Exchange, edited by N. C. Staub 8. Extrapulmonary Manifestations of Respiratory Disease, edited by E. D. Robin 9. Chronic Obstructive Pulmonary Disease, edited by T. L. Petty 10. Pathogenesis and Therapy of Lung Cancer, edited by C. C. Harris 11. Genetic Determinants of Pulmonary Disease, edited by S. D. Litwin 12. The Lung in the Transition Between Health and Disease, edited by P. T. Macklem and S. Permutt 13. Evolution of Respiratory Processes: A Comparative Approach, edited by S. C. Wood and C. Lenfant 14. Pulmonary Vascular Diseases, edited by K. M. Moser 15. Physiology and Pharmacology of the Airways, edited by J. A. Nadel 16. Diagnostic Techniques in Pulmonary Disease (in two parts), edited by M. A. Sackner 17. Regulation of Breathing (in two parts), edited by T. F. Hornbein 18. Occupational Lung Diseases: Research Approaches and Methods, edited by H. Weill and M. Turner-Warwick 19. Immunopharmacology of the Lung, edited by H. H. Newball 20. Sarcoidosis and Other Granulomatous Diseases of the Lung, edited by B. L. Fanburg
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21. Sleep and Breathing, edited by N. A. Saunders and C. E. Sullivan 22. Pneumocystis carinii Pneumonia: Pathogenesis, Diagnosis, and Treatment, edited by L. S. Young 23. Pulmonary Nuclear Medicine: Techniques in Diagnosis of Lung Disease, edited by H. L. Atkins 24. Acute Respiratory Failure, edited by W. M. Zapol and K. J. Falke 25. Gas Mixing and Distribution in the Lung, edited by L. A. Engel and M. Paiva 26. High-Frequency Ventilation in Intensive Care and During Surgery, edited by G. Carlon and W. S. Howland 27. Pulmonary Development: Transition from Intrauterine to Extrauterine Life, edited by G. H. Nelson 28. Chronic Obstructive Pulmonary Disease: Second Edition, edited by T. L. Petty 29. The Thorax (in two parts), edited by C. Roussos and P. T. Macklem 30. The Pleura in Health and Disease, edited by J. Chrétien, J. Bignon, and A. Hirsch 31. Drug Therapy for Asthma: Research and Clinical Practice, edited by J. W. Jenne and S. Murphy 32. Pulmonary Endothelium in Health and Disease, edited by U. S. Ryan 33. The Airways: Neural Control in Health and Disease, edited by M. A. Kaliner and P. J. Barnes 34. Pathophysiology and Treatment of Inhalation Injuries, edited by J. Loke 35. Respiratory Function of the Upper Airway, edited by O. P. Mathew and G. Sant’Ambrogio 36. Chronic Obstructive Pulmonary Disease: A Behavioral Perspective, edited by A. J. McSweeny and I. Grant 37. Biology of Lung Cancer: Diagnosis and Treatment, edited by S. T. Rosen, J. L. Mulshine, F. Cuttitta, and P. G. Abrams 38. Pulmonary Vascular Physiology and Pathophysiology, edited by E. K. Weir and J. T. Reeves 39. Comparative Pulmonary Physiology: Current Concepts, edited by S. C. Wood 40. Respiratory Physiology: An Analytical Approach, edited by H. K. Chang and M. Paiva 41. Lung Cell Biology, edited by D. Massaro 42. Heart–Lung Interactions in Health and Disease, edited by S. M. Scharf and S. S. Cassidy 43. Clinical Epidemiology of Chronic Obstructive Pulmonary Disease, edited by M. J. Hensley and N. A. Saunders
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44. Surgical Pathology of Lung Neoplasms, edited by A. M. Marchevsky 45. The Lung in Rheumatic Diseases, edited by G. W. Cannon and G. A. Zimmerman 46. Diagnostic Imaging of the Lung, edited by C. E. Putman 47. Models of Lung Disease: Microscopy and Structural Methods, edited by J. Gil 48. Electron Microscopy of the Lung, edited by D. E. Schraufnagel 49. Asthma: Its Pathology and Treatment, edited by M. A. Kaliner, P. J. Barnes, and C. G. A. Persson 50. Acute Respiratory Failure: Second Edition, edited by W. M. Zapol and F. Lemaire 51. Lung Disease in the Tropics, edited by O. P. Sharma 52. Exercise: Pulmonary Physiology and Pathophysiology, edited by B. J. Whipp and K. Wasserman 53. Developmental Neurobiology of Breathing, edited by G. G. Haddad and J. P. Farber 54. Mediators of Pulmonary Inflammation, edited by M. A. Bray and W. H. Anderson 55. The Airway Epithelium, edited by S. G. Farmer and D. Hay 56. Physiological Adaptations in Vertebrates: Respiration, Circulation, and Metabolism, edited by S. C. Wood, R. E. Weber, A. R. Hargens, and R. W. Millard 57. The Bronchial Circulation, edited by J. Butler 58. Lung Cancer Differentiation: Implications for Diagnosis and Treatment, edited by S. D. Bernal and P. J. Hesketh 59. Pulmonary Complications of Systemic Disease, edited by J. F. Murray 60. Lung Vascular Injury: Molecular and Cellular Response, edited by A. Johnson and T. J. Ferro 61. Cytokines of the Lung, edited by J. Kelley 62. The Mast Cell in Health and Disease, edited by M. A. Kaliner and D. D. Metcalfe 63. Pulmonary Disease in the Elderly Patient, edited by D. A. Mahler 64. Cystic Fibrosis, edited by P. B. Davis 65. Signal Transduction in Lung Cells, edited by J. S. Brody, D. M. Center, and V. A. Tkachuk 66. Tuberculosis: A Comprehensive International Approach, edited by L. B. Reichman and E. S. Hershfield 67. Pharmacology of the Respiratory Tract: Experimental and Clinical Research, edited by K. F. Chung and P. J. Barnes 68. Prevention of Respiratory Diseases, edited by A. Hirsch, M. Goldberg, J.-P. Martin, and R. Masse
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69. Pneumocystis carinii Pneumonia: Second Edition, edited by P. D. Walzer 70. Fluid and Solute Transport in the Airspaces of the Lungs, edited by R. M. Effros and H. K. Chang 71. Sleep and Breathing: Second Edition, edited by N. A. Saunders and C. E. Sullivan 72. Airway Secretion: Physiological Bases for the Control of Mucous Hypersecretion, edited by T. Takishima and S. Shimura 73. Sarcoidosis and Other Granulomatous Disorders, edited by D. G. James 74. Epidemiology of Lung Cancer, edited by J. M. Samet 75. Pulmonary Embolism, edited by M. Morpurgo 76. Sports and Exercise Medicine, edited by S. C. Wood and R. C. Roach 77. Endotoxin and the Lungs, edited by K. L. Brigham 78. The Mesothelial Cell and Mesothelioma, edited by M.-C. Jaurand and J. Bignon 79. Regulation of Breathing: Second Edition, edited by J. A. Dempsey and A. I. Pack 80. Pulmonary Fibrosis, edited by S. Hin. Phan and R. S. Thrall 81. Long-Term Oxygen Therapy: Scientific Basis and Clinical Application, edited by W. J. O’Donohue, Jr. 82. Ventral Brainstem Mechanisms and Control of Respiration and Blood Pressure, edited by C. O. Trouth, R. M. Millis, H. F. Kiwull-Schöne, and M. E. Schläfke 83. A History of Breathing Physiology, edited by D. F. Proctor 84. Surfactant Therapy for Lung Disease, edited by B. Robertson and H. W. Taeusch 85. The Thorax: Second Edition, Revised and Expanded (in three parts), edited by C. Roussos 86. Severe Asthma: Pathogenesis and Clinical Management, edited by S. J. Szefler and D. Y. M. Leung 87. Mycobacterium avium–Complex Infection: Progress in Research and Treatment, edited by J. A. Korvick and C. A. Benson 88. Alpha 1–Antitrypsin Deficiency: Biology • Pathogenesis • Clinical Manifestations • Therapy, edited by R. G. Crystal 89. Adhesion Molecules and the Lung, edited by P. A. Ward and J. C. Fantone 90. Respiratory Sensation, edited by L. Adams and A. Guz 91. Pulmonary Rehabilitation, edited by A. P. Fishman 92. Acute Respiratory Failure in Chronic Obstructive Pulmonary Disease, edited by J.-P. Derenne, W. A. Whitelaw, and T. Similowski
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93. Environmental Impact on the Airways: From Injury to Repair, edited by J. Chrétien and D. Dusser 94. Inhalation Aerosols: Physical and Biological Basis for Therapy, edited by A. J. Hickey 95. Tissue Oxygen Deprivation: From Molecular to Integrated Function, edited by G. G. Haddad and G. Lister 96. The Genetics of Asthma, edited by S. B. Liggett and D. A. Meyers 97. Inhaled Glucocorticoids in Asthma: Mechanisms and Clinical Actions, edited by R. P. Schleimer, W. W. Busse, and P. M. O’Byrne 98. Nitric Oxide and the Lung, edited by W. M. Zapol and K. D. Bloch 99. Primary Pulmonary Hypertension, edited by L. J. Rubin and S. Rich 100. Lung Growth and Development, edited by J. A. McDonald 101. Parasitic Lung Diseases, edited by A. A. F. Mahmoud 102. Lung Macrophages and Dendritic Cells in Health and Disease, edited by M. F. Lipscomb and S. W. Russell 103. Pulmonary and Cardiac Imaging, edited by C. Chiles and C. E. Putman 104. Gene Therapy for Diseases of the Lung, edited by K. L. Brigham 105. Oxygen, Gene Expression, and Cellular Function, edited by L. Biadasz Clerch and D. J. Massaro 106. Beta2-Agonists in Asthma Treatment, edited by R. Pauwels and P. M. O’Byrne 107. Inhalation Delivery of Therapeutic Peptides and Proteins, edited by A. L. Adjei and P. K. Gupta 108. Asthma in the Elderly, edited by R. A. Barbee and J. W. Bloom 109. Treatment of the Hospitalized Cystic Fibrosis Patient, edited by D. M. Orenstein and R. C. Stern 110. Asthma and Immunological Diseases in Pregnancy and Early Infancy, edited by M. Schatz, R. S. Zeiger, and H. N. Claman 111. Dyspnea, edited by D. A. Mahler 112. Proinflammatory and Antiinflammatory Peptides, edited by S. I. Said 113. Self-Management of Asthma, edited by H. Kotses and A. Harver 114. Eicosanoids, Aspirin, and Asthma, edited by A. Szczeklik, R. J. Gryglewski, and J. R. Vane 115. Fatal Asthma, edited by A. L. Sheffer 116. Pulmonary Edema, edited by M. A. Matthay and D. H. Ingbar 117. Inflammatory Mechanisms in Asthma, edited by S. T. Holgate and W. W. Busse
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118. Physiological Basis of Ventilatory Support, edited by J. J. Marini and A. S. Slutsky 119. Human Immunodeficiency Virus and the Lung, edited by M. J. Rosen and J. M. Beck 120. Five-Lipoxygenase Products in Asthma, edited by J. M. Drazen, S.-E. Dahlén, and T. H. Lee 121. Complexity in Structure and Function of the Lung, edited by M. P. Hlastala and H. T. Robertson 122. Biology of Lung Cancer, edited by M. A. Kane and P. A. Bunn, Jr. 123. Rhinitis: Mechanisms and Management, edited by R. M. Naclerio, S. R. Durham, and N. Mygind 124. Lung Tumors: Fundamental Biology and Clinical Management, edited by C. Brambilla and E. Brambilla 125. Interleukin-5: From Molecule to Drug Target for Asthma, edited by C. J. Sanderson 126. Pediatric Asthma, edited by S. Murphy and H. W. Kelly 127. Viral Infections of the Respiratory Tract, edited by R. Dolin and P. F. Wright 128. Air Pollutants and the Respiratory Tract, edited by D. L. Swift and W. M. Foster 129. Gastroesophageal Reflux Disease and Airway Disease, edited by M. R. Stein 130. Exercise-Induced Asthma, edited by E. R. McFadden, Jr. 131. LAM and Other Diseases Characterized by Smooth Muscle Proliferation, edited by J. Moss 132. The Lung at Depth, edited by C. E. G. Lundgren and J. N. Miller 133. Regulation of Sleep and Circadian Rhythms, edited by F. W. Turek and P. C. Zee 134. Anticholinergic Agents in the Upper and Lower Airways, edited by S. L. Spector 135. Control of Breathing in Health and Disease, edited by M. D. Altose and Y. Kawakami 136. Immunotherapy in Asthma, edited by J. Bousquet and H. Yssel 137. Chronic Lung Disease in Early Infancy, edited by R. D. Bland and J. J. Coalson 138. Asthma’s Impact on Society: The Social and Economic Burden, edited by K. B. Weiss, A. S. Buist, and S. D. Sullivan 139. New and Exploratory Therapeutic Agents for Asthma, edited by M. Yeadon and Z. Diamant 140. Multimodality Treatment of Lung Cancer, edited by A. T. Skarin
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141. Cytokines in Pulmonary Disease: Infection and Inflammation, edited by S. Nelson and T. R. Martin 142. Diagnostic Pulmonary Pathology, edited by P. T. Cagle 143. Particle–Lung Interactions, edited by P. Gehr and J. Heyder 144. Tuberculosis: A Comprehensive International Approach, Second Edition, Revised and Expanded, edited by L. B. Reichman and E. S. Hershfield 145. Combination Therapy for Asthma and Chronic Obstructive Pulmonary Disease, edited by R. J. Martin and M. Kraft 146. Sleep Apnea: Implications in Cardiovascular and Cerebrovascular Disease, edited by T. D. Bradley and J. S. Floras 147. Sleep and Breathing in Children: A Developmental Approach, edited by G. M. Loughlin, J. L. Carroll, and C. L. Marcus 148. Pulmonary and Peripheral Gas Exchange in Health and Disease, edited by J. Roca, R. Rodriguez-Roisen, and P. D. Wagner 149. Lung Surfactants: Basic Science and Clinical Applications, R. H. Notter 150. Nosocomial Pneumonia, edited by W. R. Jarvis 151. Fetal Origins of Cardiovascular and Lung Disease, edited by David J. P. Barker 152. Long-Term Mechanical Ventilation, edited by N. S. Hill 153. Environmental Asthma, edited by R. K. Bush 154. Asthma and Respiratory Infections, edited by D. P. Skoner 155. Airway Remodeling, edited by P. H. Howarth, J. W. Wilson, J. Bousquet, S. Rak, and R. A. Pauwels 156. Genetic Models in Cardiorespiratory Biology, edited by G. G. Haddad and T. Xu 157. Respiratory-Circulatory Interactions in Health and Disease, edited by S. M. Scharf, M. R. Pinsky, and S. Magder 158. Ventilator Management Strategies for Critical Care, edited by N. S. Hill and M. M. Levy 159. Severe Asthma: Pathogenesis and Clinical Management, Second Edition, Revised and Expanded, edited by S. J. Szefler and D. Y. M. Leung 160. Gravity and the Lung: Lessons from Microgravity, edited by G. K. Prisk, M. Paiva, and J. B. West 161. High Altitude: An Exploration of Human Adaptation, edited by T. F. Hornbein and R. B. Schoene 162. Drug Delivery to the Lung, edited by H. Bisgaard, C. O’Callaghan, and G. C. Smaldone 163. Inhaled Steroids in Asthma: Optimizing Effects in the Airways, edited by R. P. Schleimer, P. M. O’Byrne, S. J. Szefler, and R. Brattsand
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164. IgE and Anti-IgE Therapy in Asthma and Allergic Disease, edited by R. B. Fick, Jr., and P. M. Jardieu 165. Clinical Management of Chronic Obstructive Pulmonary Disease, edited by T. Similowski, W. A. Whitelaw, and J.-P. Derenne 166. Sleep Apnea: Pathogenesis, Diagnosis, and Treatment, edited by A. I. Pack 167. Biotherapeutic Approaches to Asthma, edited by J. Agosti and A. L. Sheffer 168. Proteoglycans in Lung Disease, edited by H. G. Garg, P. J. Roughley, and C. A. Hales 169. Gene Therapy in Lung Disease, edited by S. M. Albelda 170. Disease Markers in Exhaled Breath, edited by N. Marczin, S. A. Kharitonov, M. H. Yacoub, and P. J. Barnes 171. Sleep-Related Breathing Disorders: Experimental Models and Therapeutic Potential, edited by D. W. Carley and M. Radulovacki 172. Chemokines in the Lung, edited by R. M. Strieter, S. L. Kunkel, and T. J. Standiford 173. Respiratory Control and Disorders in the Newborn, edited by O. P. Mathew 174. The Immunological Basis of Asthma, edited by B. N. Lambrecht, H. C. Hoogsteden, and Z. Diamant 175. Oxygen Sensing: Responses and Adaptation to Hypoxia, edited by S. Lahiri, G. L. Semenza, and N. R. Prabhakar 176. Non-Neoplastic Advanced Lung Disease, edited by J. R. Maurer 177. Therapeutic Targets in Airway Inflammation, edited by N. T. Eissa and D. P. Huston 178. Respiratory Infections in Allergy and Asthma, edited by S. L. Johnston and N. G. Papadopoulos 179. Acute Respiratory Distress Syndrome, edited by M. A. Matthay 180. Venous Thromboembolism, edited by J. E. Dalen 181. Upper and Lower Respiratory Disease, edited by J. Corren, A. Togias, and J. Bousquet 182. Pharmacotherapy in Chronic Obstructive Pulmonary Disease, edited by B. R. Celli 183. Acute Exacerbations of Chronic Obstructive Pulmonary Disease, edited by N. M. Siafakas, N. R. Anthonisen, and D. Georgopoulos 184. Lung Volume Reduction Surgery for Emphysema, edited by H. E. Fessler, J. J. Reilly, Jr., and D. J. Sugarbaker 185. Idiopathic Pulmonary Fibrosis, edited by J. P. Lynch III 186. Pleural Disease, edited by D. Bouros
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187. Oxygen/Nitrogen Radicals: Lung Injury and Disease, edited by V. Vallyathan, V. Castranova, and X. Shi 188. Therapy for Mucus-Clearance Disorders, edited by B. K. Rubin and C. P. van der Schans 189. Interventional Pulmonary Medicine, edited by J. F. Beamis, Jr., P. N. Mathur, and A. C. Mehta 190. Lung Development and Regeneration, edited by D. J. Massaro, G. Massaro, and P. Chambon 191. Long-Term Intervention in Chronic Obstructive Pulmonary Disease, edited by R. Pauwels, D. S. Postma, and S. T. Weiss 192. Sleep Deprivation: Basic Science, Physiology, and Behavior, edited by Clete A. Kushida 193. Sleep Deprivation: Clinical Issues, Pharmacology, and Sleep Loss Effects, edited by Clete A. Kushida 194. Pneumocystis Pneumonia: Third Edition, Revised and Expanded, edited by P. D. Walzer and M. Cushion 195. Asthma Prevention, edited by William W. Busse and Robert F. Lemanske, Jr. 196. Lung Injury: Mechanisms, Pathophysiology, and Therapy, edited by Robert H. Notter, Jacob Finkelstein, and Bruce Holm 197. Ion Channels in the Pulmonary Vasculature, edited by Jason X.-J. Yuan 198. Chronic Obstuctive Pulmonary Disease: Cellular and Molecular Mechanisms, edited by Peter J. Barnes 199. Pediatric Nasal and Sinus Disorders, edited by Tania Sih and Peter A. R. Clement 200. Functional Lung Imaging, edited by David Lipson and Edwin van Beek 201. Lung Surfactant Function and Disorder, edited by Kaushik Nag 202. Pharmacology and Pathophysiology of the Control of Breathing, edited by Denham S. Ward, Albert Dahan and Luc J. Teppema 203. Molecular Imaging of the Lungs, edited by Daniel Schuster and Timothy Blackwell 204. Air Pollutants and the Respiratory Tract: Second Edition, edited by W. Michael Foster and Daniel L. Costa
The opinions expressed in these volumes do not necessarily represent the views of the National Institutes of Health.
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PHARMACOLOGY AND PATHOPHYSIOLOGY OF THE CONTROL OF BREATHING
Edited by
Denham S. Ward University of Rochester Medical Center Rochester, New York, U.S.A.
Albert Dahan Leiden University Medical Center Leiden, Netherlands
Luc J. Teppema Leiden University Medical Center Leiden, Netherlands
Boca Raton London New York Singapore
DK1337_Discl.fm Page 1 Wednesday, April 13, 2005 3:16 PM
Published in 2005 by Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2005 by Taylor & Francis Group, LLC No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-10: 0-8247-5890-0 (Hardcover) International Standard Book Number-13: 978-0-8247-5890-5 (Hardcover) Library of Congress Card Number 2005043947 This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the consequences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe.
Library of Congress Cataloging-in-Publication Data Ward, Denham. Pharmacology and pathophysiology of the control of breathing / Denham Ward. p. cm. Includes bibliographical references and index. ISBN 0-8247-5890-0 1. Respiration--Regulation. 2. Lungs--Pathophysiology. 3. Pulmonary pharmacology. 4. Inhalation anesthesia. I. Title. QP123.W37 2005 612.2--dc22
2005043947
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INTRODUCTION
‘‘What is the use of breathing? That it is not a trifling use is clear from our inability to survive for even the shortest time after it has stopped. Hence also it is obvious that its importance is not for any particular and partial activity, but for life itself.’’ De Usu Respirationis Galen (c.120–c.200) In a second writing titled De Causis Respirationis, Galen went on to say: ‘‘It is impossible either to confirm the hypothesis of breathing or to put it right if it is impeded or completely stopped, without knowing its causes.’’ Galen’s issues are, in fact, the subject of this latest addition to the series Lung Biology in Health and Disease—Pharmacology and Pathophysiology of the Control of Breathing. This topic has fascinated generations of fundamental and clinical researchers, and their contributions have played an essential role in our current understanding of the control of breathing in health and diseases. And today the names of many of these scientific giants grace the virtual walls of the Biomedicine Hall of Fame! A turning point in the evolution of this work was the presentation of the Silliman Lectures given in 1916 by J.S. Haldane at Yale University. That was followed in 1921 by the publication of Respiration by J.S. Haldane and J.G. Priestley. In the preface to this monumental book, the authors acknowledged that ‘‘many gaps remain to be filled’’ but ‘‘the observations and experiments required [to fill them] are not yet available.’’ Today, almost a century later, many of the gaps have been filled thanks to the brilliant and dedicated researchers who have worked in this field. Nonetheless, the road to understanding the control of breathing has vii
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been long and tortuous. First, researchers observed changes in breathing patterns that occur with environmental variations, especially altitude, in an effort to uncover the mechanisms of breathing control. Then, they turned attention to the impact of pathology on this process, and breathing disruptions caused by disease became a fertile topic of investigation. In many ways, the 15 volumes on (or related to) control of breathing that we have published illustrate the purpose of this series when it was conceived more than three decades ago — namely, to report about and stimulate areas of biology and medicine, especially those that are so amazingly dynamic. This volume takes us to a new level by filling many of the gaps that Haldane and Priestley identified. In addition, as Thomas Hornbein says in his Foreword, ‘‘Within this volume are many enticing next steps.’’ The series of monographs Lung Biology in Health and Disease is most pleased to present this volume to its readership and I, personally, owe a great debt of gratitude to the editors, Drs. Denham S. Ward, Albert Dahan, and Luc Teppema, and the many authors for the privilege of introducing this important new contribution. Claude Lenfant, MD Gaithersburg, Maryland
PREFACE
We would be seriously remiss if we did not first acknowledge the work of the ‘‘fourth editor’’ of this volume, Debra L. Lipscomb. Without her careful scrutiny of every chapter and her abilities to pick out problems in meaning, grammar, spelling and references, the quality of this volume would have been severely compromised. Previous volumes in this series have dealt with the regulation of breathing and it was the volume edited by Dr. Thomas Hornbein in 1981 that has served as an essential reference for us during our careers. We are particularly gratified that he has provided a Foreword to this volume. Subsequent volumes in this series dealing with the control of breathing, as well as the two volumes in the American Physiological Society’s Handbook of Physiology have provided up-to-date summaries of the field in the face of a rapid increase in knowledge. Pharmacology has long played an important role in the control of breathing, both in providing tools for physiological experiments and in understanding the ventilatory effects of pharmacological agents. The increase in knowledge in both physiology and pharmacology has greatly extended our knowledge of the cellular and subcellular elements involved in controlling ventilation. Drug effects on control of breathing were covered as a single chapter in both Hornbein’s original volume as well as in the Handbook of Physiology. However, the increase in our understanding of the mechanisms of action of pharmacological agents on the control of breathing now warrants a separate volume. We have attempted to organize this knowledge by first reviewing the relevant physiology from a perspective of the substrate for pharmacological action and also by reviewing pharmacology principles as they can be applied to the control of breathing. We then have selected topics ix
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of pathophysiology and pharmacology that are relevant to clinical practice. It is hoped that this organization, rather than organizing by drug, will provide a more useful reference for both clinicians and research scientists. There are many clinical problems both in respiratory side effects of drugs and in finding drugs that will treat abnormalities in the control of breathing that remain to be solved. The increased understanding of the specific ion channels, receptors and neurotransmitters involved in respiratory control provides important information for pharmacological research. We hope this volume will set the stage for future research and provide as much motivation for beginning reseachers as Dr. Hornbein’s volume did for us.
FOREWORD
When I was young I fell in love with the carotid body. Though I did not know it at the time, this infatuation was kindled when my prepubertal proclivity for climbing things—trees and houses—bumped into a teenage discovery: mountains were better. With medical school my addiction to the reading of mountaineering literature was supplanted by exploration of the literature about human adaptation to high altitude. One paper, in particular, became seminal. I found in a report by Hugo Chiodi, a Peruvian clinician-investigator, that when permanent residents of high altitude were compared to lowlanders acclimatized for a time to the same altitude, the highlanders appeared to exhibit less ventilation (higher alveolar PCO2 ) and greater polycythemia than the lowlanders [1]. Was there a connection, I wondered, between the two? Did the hypoventilation provoke the greater polycythemia? Or might it be the other way around, that polycythemia somehow enables less ventilation? Or might both possibilities coexist in a kind of positive feedback system? During my final year in medical school, I seized upon a six-week elective period to embark upon my first research project, to study the effect of polycythemia on breathing. With the oversight of Albert Roos, who would become the mentor for my subsequent research training, I transfused myself with five units of blood to elevate my hematocrit from 45% to 60%. Pedaling away on a cycle ergometer while breathing oxygen mixtures to simulate different altitudes, Dr. Roos and I compared my breathing at high hematocrit with that at normal hematocrit. Not surprisingly, ventilation was less at the higher hematocrit. I cannot say it was significantly less, for this paper, ‘Effect of Polycythemia on Respiration’ was published in spite of being performed on only one subject (other volunteers were hard to find) and hence had no statistical analysis [2]. xi
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Foreword
Next came the question, how does polycythemia result in less exercise hyperpnea? Enter the carotid body and the beginnings of an affair with chemoreception and the chemical control of breathing that directed my scientific exploration for the next quarter century. At the time I climbed Mount Everest in 1963 [3], I never imagined the numbers that would be thronging to its top four decades later. Nor did I, the first time I hung the Nerve of Hering across a pair of platinum electrodes and listened to the crackling crescendo of neuronal traffic as the animal’s PO2 was lowered, anticipate the current profundity of inquiry into the ventilatory control system. The pharmacology of ventilatory control comprised a single chapter in the original volume on regulation of breathing in this series [4], as well as its second edition [5]; that pharmacology could command an entire, rich volume of its own was unimaginable not so long ago. This book (number 202 In Claude Lenfant’s series, Lung Biology in Health and Disease), Pharmacology and Pathophysiology of the Control of Breathing, has been creatively conceived and gestated by Drs. Ward, Dahan, and Teppema. The book is divided into three parts: Neuropharmacology and Physiology, Pathophysiology, and Clinical Pharmacology. The first section is, for me, the dessert, even though it comes at the beginning of the meal. This section explores the functional anatomy and physiology of the ventilatory control system from the intracellular to the integrative level, including what is coming to be known about the roles of nervous system plasticity and genes. It’s like the woods out back where one could go to explore, full of intrigue and an inexhaustible supply of new questions emerging from every answer. This first section stirred past wonderings of a time when I puzzled about why inhalational anesthetics, all of which produced anesthesia, differed in the fingerprint each left on the central nervous system with regard to side effects such as breathing. For example, diethyl ether, the founding father of general anesthesia, produced little or no ventilatory depression, as measured by the alveolar or arterial PCO2, until anesthesia became quite deep. This preservation seemed to result from an increasing tachypnea sufficient to offset the progressive diminution of tidal volume. Most other anesthetics were associated with dose-related diminutions in alveolar ventilation, some much more than others. Each anesthetic seemed to have a ventilatory identity of its own with regard to such dose-related parameters as tidal volume, ventilatory drive (Vt /Ti), relative timing of inspiration and expiration, frequency, and even the nature of inspiratory and expiratory pauses. How do we explain such differences? Do different drugs act upon different cells, or different receptors, or in different ways to account for their unique signatures? Could understanding where and how they work provide clues about how the various components of the ventilatory control
Foreword
xiii
system are put together and how the separate parts communicate with each other under physiological as well as pharmacological circumstances? My wonderings never became more than that. But one need only reflect on the potency of the tools currently available to our imaginations to realize there is world of understanding out there waiting to be explored. Within this volume are many enticing next steps. Thomas Hornbein, M.D. Professor Emeritus Departments of Anesthesiology and Physiology and Biophysics University of Washington Seattle, Washington
References 1. 2. 3. 4.
5.
Chiodi, H., Respiratory adaptations to chronic high altitude hypoxia, J. Appl. Physiol. 10, 81–87, 1957. Hornbein, T.F. and Roos, A., Effect of polycythemia on respirations, J. Appl. Physiol. 12, 86–90, 1958. Hornbein, T.F., Everest, the West Ridge, San Francisco, The Sierra Club, 1965. Hornbein, T.F., ed., Regulation of Breathing, New York, Marcel Dekker, Inc., 17, 1436 pp., 1981 (One of a series of monographs in Lung Biology in Health and Disease, edited by Lenfant, C.) Dempsey, J.D. and Pack, A., eds., Regulation of Breathing, Second edition, New York, Marcel Dekker, Inc., 79, 1219 pp., 1995 (One of a series of mongraphs in Lung Biology in Health and Disease, edited by Lenfant, C.)
CONTRIBUTORS
Khalid F. Almoosa, M.D., F.C.C.P. Clinical Instructor of Medicine, University of Cincinnati College of Medicine, Cincinnati, Ohio. Peter L. Bailey, M.D. Professor, Department of Anesthesiology, University of Rochester School of Medicine and Dentistry, Rochester, New York. Ryan W. Bavis, Ph.D. Assistant Professor, Department of Biology, Bates College, Lewiston, Maine. Philip E. Bickler, M.D. Professor, Department of Anesthesiology, University of California, San Francisco, California. Dante A. Cerza University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania. Albert Dahan, M.D., Ph.D. Professor, Department of Anesthesiology, Leiden University Medical Center, Leiden, The Netherlands. Lars I. Eriksson, M.D. Professor and Academic Chair, Department of Anesthesiology, Karolinska Hospital and Institute, Stockholm, Sweden. David D. Fuller, Ph.D. Assistant Professor, Department of Physical Therapy University of Florida, Gainesville, Florida. David Gozal, M.D. Professor and Children’s Foundation Chair for Pediatric Research, Vice Chair for Research, Director, Kosair Children’s Hospital Research Institute, Department of Pediatrics University of Louisville Louisville, Kentucky.
xv
xvi
Contributors
Jeffrey B. Gross, M.D. Professor, Departments of Anesthesiology, University of Connecticut School of Medicine Farmington, Connecticut. Shiroh Isono, M.D. Department of Anesthesiology, Chiba University Graduate School of Medicine, Chiba, Japan Shahrokh Javaheri, M.D. Professor, Department of Medicine, University of Cincinnati College of Medicine Cincinnati, Ohio and SleepCare Diagnostics Mason, Ohio. Suzanne Karan, M.D. Assistant Professor, Department of Anesthesiology University of Rochester School of Medicine and Dentistry, Rochester, New York. Gordon S. Mitchell, Ph.D. Department of Comparative Biosciences, University of Wisconsin, School of Veterinary Medicine, Madison, Wisconsin. Shakeeb H. Moosavi, Ph.D. Harvard School of Public Health, Boston, Massachusetts, and Imperial College School of Medicine, London, England. Takashi Nishino, M.D. Professor and Chair, Department of Anesthesiology, Chiba University Graduate School of Medicine, Chiba, Japan. Colin A. Nurse, Ph.D. Department of Biology, McMaster University, Hamilton, Ontario, Canada. Erik Olofsen, M.S. Department of Anesthesiology, Leiden University Medical Center, Leiden, The Netherlands. Susheel P. Patil, Ph.D. Instructor, Division of Pulmonary and Critical Care Medicine Department of Medicine, Johns Hopkins University Baltimore, Maryland. David Paydarfar, M.D. Associate Professor of Neurology and Physiology, Department of Neurology, University of Massachusetts, Medical School Worcester, Massachusetts. Frank L. Powell, Ph.D. Professor, Department of Medicine, Director, White Mountain Research Station, University of California, San Diego, La Jolla, California. Raymonda Romberg, M.D., Ph.D. Department of Anesthesiology, Leiden University Medical Center, Leiden, The Netherlands. Jacob Rosenberg, M.D.
University of Copenhagen, Hellerup, Denmark.
Contributors
xvii
Elise Sarton, M.D. Department of Anesthesiology, Leiden University Medical Center, Leiden, The Netherlands. Hartmut Schneider, M.D., Ph.D. Department of Medicine and Division of Pulmonary and Critical Care Medicine, Johns Hopkins University, Baltimore, Maryland. Alan R. Schwartz, M.D. Department of Medicine and Division of Pulmonary and Critical Care Medicine, Johns Hopkins University, Baltimore, Maryland. Steven A. Shea, Ph.D. Associate Professor, Department of Medicine, Harvard Medical School, and Director, Medical Chronobiology Program, Brigham and Women’s Hospital, Boston, Massachusetts. Philip L. Smith Sleep Disorders Center, Johns Hopkins University Baltimore, Maryland. Kingman P. Strohl, M.D. Professor, Departments of Medicine, Human Genetics, and Epidemiology and Biostatistics, Case Western Reserve University, Cleveland, Ohio. Astrid G. Stucke, M.D. Research Fellow, Department of Anesthesiology, Medical College of Wisconsin, Milwaukee, Wisconsin. Eckehard A.E. Stuth, M.D. Associate Professor, Department of Anesthesiology, Medical College of Wisconsin, and Director, Cardiac Anesthesia, Children’s Hospital of Wisconsin, Milwaukee, Wisconsin. Luc J. Teppema, Ph.D. Department of Anesthesiology, Leiden University Medical Center, Leiden, The Netherlands. Rajbala Thakur, M.D. Associate Professor, Department of Anesthesiology, University of Rochester School of Medicine and Dentistry, Rochester, New York. Denham S. Ward, M.D., Ph.D. Professor, Departments of Anesthesiology and Biomedical Engineering, University of Rochester Medical Center, Rochester, New York. David O. Warner, M.D. Professor, Department of Anesthesiology, Mayo Medical School and Mayo Clinic Rochester, Rochester, Minnesota. Edward J. Zuperku, Ph.D. Research Professor, Department of Anesthesiology, Medical College of Wisconsin, Milwaukee, Wisconsin.
ABBREVIATIONS
ABBREVIATION
DEFINITION
5-HT 5-HTP ACE ACh ACLS AG II AHI AHR AMPA
serotonin 5-hydroxytryptophan angiotensin converting enzyme acetylcholine advanced cardiac life support angiotensin II apnea-hypopnea index acute hypoxic response a-amino-3-hydroxy-5-methylisoxazole4-propionate acute mountain sickness atrial naturetic peptide area postrema action potential 2-amino-5phosphonovalerat artificial brainstem perfusion apolipoprotein E adenosine triphosphate arginine vasopressin blood-brain barrier brain blood flow bilevel positive airway pressure bispectral index large conductance Kþ body mass index blood pressure
AMS ANP AP AP AP5 APB APOE ATP AVP BBB BBF BiPAP BIS BK BMI BP
xix
xx BPM BSA CA CA CAD CB CBD CCHS CCK CDR CH CIH CMS CNS COPD CPAP CPG CPR CRP CSF CSN CVLM CVMS CVO CVRG D2-R DA DEF DI DLCO DM DRG DZ E EBSN EC50 ECE-1 ECF EDIA EMG EPI EPSP ERV FEF
Abbreviations breaths per minute body surface area carbonic anhydrase catecholamine coronary artery disease carotid body carotid body denervation congenital central hypoventilation syndrome cholecystokinin chronic dorsal rhizotomy chronic hypoxia chronic intermittent hypoxia chronic mountain sickness central nervous system chronic obstructive pulmonary disease continuous positive airway pressure central pattern generator cardio-pulmonary resuscitation C-reactive protein cerebrospinal fluid carotid sinus nerve caudal ventrolateral medulla caudal ventral medullary surface circumventricular organs caudal ventral respiratory group D2 receptor dopamine dynamic end-tidal forcing diaphragm single breath diffusion capacity dorsomedial hypothalamus dorsal respiratory group dizygotic expiratory expiratory bulbospinal neuron concentration yielding 50% effect endothelin-converting enzyme-1 extracellular fluid EMG activity in the diaphragm electromyelogram epinephrine excitatory postsynaptic potential expiratory reserve volume forced expiratory flow
Abbreviations fMRI fR FRC FVC GABA GER GG GiA GPN HAPE HCVR HD HEPES HERG HIF HM HPVR HR HTN HVA HVD HVR I IBSN IBW IC IH IM ION IPPB IPSP IT IV Kca Kir KO LC LHB LPB LPG LRN LTF LV LVA
xxi functional magnetic resonance imaging respiratory frequency functional residual capacity forced vital capacity g-aminobutyric acid gastroesophageal reflux genioglossus gigantocellular nucleus pars a glossopharyngeal nerve high-altitude pulmonary edema hypercapnic ventilatory response hypoxic desensitization 2-hydroxyethyl-l-piperazineethane sulfonic acid human ether-a-gogo-related gene hypoxic-inducible factor hypoglossal motoneuron hypoxic pulmonary vasoconstrictor response heart rate hypertension high-voltage activated hypoxic ventilatory depression hypoxic ventilatory response inspiratory inspiratory bulbospinal neuron ideal body weight intercostal intermittent hypoxia intramuscular infraorbital nerve intermittent positive-pressure breathing inhibitory postsynaptic potential intrathecal intravenous Ca2þ-activated Kþ channel inward rectifying Kþ channel knock-out locus coeruleus lateral habenular nucleus lateral parabrachial nucleus lateral paragigantocellular nucleus lateral reticular nucleus long-term facilitation left ventricle low-voltage activated
xxii LVH MAC MAP MFBS MI MP MRI MS MVV MZ NA nACh receptor nAChR NADP NBQX NE NEB NEP NEPI NF NHE NIMV NMBA NMDA NO NOS NOS-1 NPY NREM NTS ODI OHS OSA OSAH OSAS OT P PACU PAG PB PBW PCA PCEA
Abbreviations left ventricular hypertrophy minimum alveolar concentration mean arterial pressure multi-frequency binary sequence myocardial infarction membrane potential magnetic resonance imaging morphine sulfate maximum voluntary ventilation monozygotic nucleus ambiguus nicotinic acetylcholine receptor nicotinic acetylcholine receptor nicotinamide adenine dinucleotide phosphate 2,3-dihydroxy-6-nitro-7-sulfamoylbenzo(f )quinoxaline norepinephrine neuroepithelial body neutral endopeptidase norepinephrine neurofilament Naþ/Hþ exchanger non-invasive mechanical ventilation neuromuscular blocking agent N-methyl D-aspartate nitric oxide nitric oxide synthase nitric oxide synthase-1 neuropeptide Y non-rapid eye movement nucleus tractus solitarius oxygen desaturation index obesity hypoventilation syndrome obstructive sleep apnea obstructive sleep apnea-hypopnea syndrome obstructive sleep apnea syndrome optic tract pyramidal tract postanesthesia care unit periaqueductal gray matter periodic breathing predicted body weight patient-controlled analgesia patient-controlled epidural analgesia
Abbreviations p-CPA Pcrit PDGF-b PEEP PEFR PEMAX PET PG PGCI pHe PHFD pHi PIIA PIIMAX PKC PK-PD PN PNDS PNG PNN POMC PPL pre-Bo¨tc PSR RAR rCBF REM RM RN ROS ROS RP RTN RV RVLM RVMM rVMS SAR SCI SD SDB SDH SIDS SK
xxiii para-chlorophenylalanine critical pressure platelet-derived growth factor b positive end-expiratory pressure peak expiratory flow rate maximum expiratory pressure positron emission tomography petrosal ganglion paragigantocellular nucleus extracellular pH post-hypoxia frequency decline intracellular pH post-inspiratory inspiratory activity maximum inspiratory pressure protein kinase C pharmacokinetic-pharmacodynamic nasal pressure post-nasal drip syndrome phrenic neurogram posterior nasal nerve proopiomelanocortin pleural pressure pre-Bo¨tziger complex pulmonary stretch receptor rapidly adapting receptor regional cerebral blood flow rapid eye movement raphe magnus raphe nuclei radical oxygen species reactive oxygen species raphe pallidus retrotrapezoid nucleus residual volume rostral ventral lateral medulla rostral ventral medial medulla rostral ventral medullary surface slowly adapting receptor spinal cord injury Sprague-Dawley strain of rats sleep-disordered breathing succinate dehydrogenase sudden infant death syndrome small-conductance Kþ channel
xxiv SLN SMA SNO SON SP STD sTEA STP SWS TASK TE TH TI TIA TLC TMP TMS TRH TTX TV UA UARS VAH VC VCO2 VDH VEGF vIPAG VMH VO2 VRG VT
Abbreviations superior laryngeal nerve supplementary motor area S-nitrosothiol supraoptic nucleus substance P short-term depression segmental high thoracic epidural anesthesia short-term potentiation slow wave sleep tandem pore acid sensitive Kþ channel expiratory phase duration tyrosine hydroxylase inspiratory phase duration transient ischemic attack total lung capacity transmural pressure transcranial magnetic stimulation thyrotropin-releasing hormone tetrodotoxin tidal volume upper airway upper airway resistance syndrome ventilatory acclimatization to hypoxia vital capacity CO2 production ventilatory deacclimatization from hypoxia vascular endothelial growth factor parabrachial nucleus ventromedial hypothalamus oxygen consumption ventral respiratory group tidal volume
CONTENTS
Introduction Preface Foreword Contributors Abbreviations
Claude Lenfant Thomas L. Hornbein
vii ix xi xv xix
I.
NEUROPHARMACOLOGY AND PHYSIOLOGY
1
1.
Peripheral Chemoreceptors: Sensors of Metabolic Status Colin A. Nurse
3
I. II.
3
Introduction Cellular Organization and Innervation of the Carotid Body III. Carotid Body Chemoreceptors: O2-Sensitive Kþ Channels IV. What is the O2 Sensor and How is it Linked to Kþ Channel(s)? V. Role of Fast-Acting Neurotransmitters in Chemosensory Processing VI. Glucose Sensing in the Carotid Body VII. Neuromodulation in the Carotid Body VIII. Future Directions Acknowledgments References 2.
4 6 8 8 12 12 15 15 16
Central Chemoreceptors Luc J. Teppema and Albert Dahan
21
I.
21
Introduction
xxv
xxvi
Contents II. III. IV.
3.
Location of Central Chemoreceptors Mechanism of Central Chemoreception Central Chemoreceptors and Breathing References
Suprapontine Control of Breathing Shakeeb H. Moosavi, David Paydarfar, and Steven A. Shea
71
I. II. III. IV. V. VI.
71 72 72 75 84
Introduction Definitions and Terminology Volitional Control Involuntary Emotional Influences Tonic Excitatory and Inhibitory Drives Interaction between ‘Behavioral’ and ‘Automatic’ Control VII. Learned Respiratory Behaviors VIII. Summary Acknowledgments References 4.
5.
24 38 47 55
86 88 90 91 91
Measurement of Drug Effects on Ventilatory Control Denham S. Ward
103
I. II. III. IV. V. VI. VII. VIII.
103 106 108 109 110 117 120 122 124
Introduction Measurement Techniques Quantification of Drug Pharmacodynamics Resting Measurements Hypercapnic Ventilatory Response Hypoxic Ventilatory Response Changes in Airway Pressure Other Stimuli References
Response Surface Modeling of Drug Interactions: Model Selection and Multimodel Inference Using the Bootstrap Erik Olofsen and Albert Dahan I. II. III. IV. V. VI.
Introduction Pharmacodynamic Interaction Models Model Selection and Multimodel Inference The Bootstrap Applications Conclusions References
133 133 134 137 140 143 151 152
Contents 6.
Respiratory Neuroplasticity: Respiratory Gases, Development, and Spinal Injury David D. Fuller, Gordon S. Mitchell, and Ryan W. Bavis I. II. III. IV. V. VI.
7.
xxvii
Introduction Plasticity Induced by Respiratory Gases in Adult Mammals Developmental Plasticity and the Control of Breathing Sex Hormones Spinal Cord Injury (SCI) and Respiratory Plasticity Conclusion Acknowledgments References
155 158 174 187 187 198 199 199
Airway Reflexes in Humans Takashi Nishino
225
I. II. III.
225 226
Introduction Central Nervous System Afferent Innervation of the Upper Airway and Receptors IV. Reflex Responses from the Upper Airway V. Afferent Innervation of the Lower Airway and Receptors VI. Integrative Aspects of the Airway Reflexes Elicited from the Lower Airways VII. Clinical Problems Associated with Airway Reflexes VIII. Conclusions Acknowledgments References 8.
155
226 229 234 238 244 250 251 251
Inheritance and Ventilatory Behavior in Animal Models Kingman P. Strohl
261
I. II.
262
III. IV. V. VI. VII.
Introduction Evidence and Implications for Inheritance of Ventilatory Traits in Humans Targeting Ventilatory Traits in Small Animals Evidence for the Inheritance of Ventilatory Traits in Rodents Estimates of the Strength of Inheritance Studies of Gene Effects in Rodent Models A Physiogenetic Map of Ventilatory Behavior
262 266 271 274 276 279
xxviii
Contents VIII. Overview and Future Directions Acknowledgments References
282 283 283
II.
PATHOPHYSIOLOGY
293
9.
Congenital Central Hypoventilation Syndrome: Should We Rename it Congenital Autonomopathy? David Gozal
295
I. II. III. IV. V.
10.
11.
Introduction Definition and Diagnosis Pathophysiology Animal Models Structural Central Nervous System Abnormalities VI. Physiologic Abnormalities of Ventilatory Control VII. Autonomic Nervous System Dysfunction VIII. Summary and Conclusions Acknowledgment References
295 296 298 300
Upper Airway Obstruction in Sleep Apnea Susheel P. Patil, Hartmut Schneider, Philip L. Smith, and Alan R. Schwartz
313
I. II. III. IV. V.
313 314 317 333 335 336
Introduction Epidemiologic and Clinical Risk Factors Pathogenesis of Upper Airway Obstruction Therapeutic Implications Summary and Conclusions References
300 301 304 304 304 304
High Altitude Frank L. Powell and Philip E. Bickler
357
I. II. III. IV.
357 358 360
V.
Introduction Ventilatory Response to High Altitude Time Domains of the HVR Increases in the Hypercapnic Ventilatory Response (HCVR) with Acclimatization High Altitude Diseases and Ventilatory Control References
372 372 376
Contents 12.
xxix
Obesity and the Control of Breathing Khalid F. Almoosa and Shahrokh Javaheri
383
I. II. III. IV.
383 384 388
V. VI.
Introduction Overview of Obesity Effects of Obesity on the Respiratory System Disorders of Ventilatory Control Associated with Obesity Effects of Treatment of Obesity and OSAH on Ventilatory Control and PaCO2 Conclusion References
399 408 412 412
III.
CLINICAL PHARMACOLOGY
423
13.
Pain Management and Regional Anesthesia Peter L. Bailey and Rajbala Thakur
425
I. II. III. IV. V.
425 426 433 438
Introduction Postoperative Respiratory Dysfunction Other Effects of Pain and Surgical Trauma Pain Control and Chronic Pain Effects of Pain and Pain Management on Respiratory Function VI. Regional Analgesia VII. Other Analgesic Agents VIII. Pre-Emptive Analgesia IX. Perioperative Analgesia and Pulmonary Outcome X. Cardiac Surgery XI. Summary References 14.
Ventilatory Effects of Medications Used for Moderate and Deep Sedation Jeffrey B. Gross and Dante A. Cerza I. II. III. IV. V. VI.
Introduction Sedatives Opioids Drug Combinations Strategies for Minimizing Respiratory Risks of Sedation Conclusion References
438 454 473 483 483 487 489 490
513 513 516 540 553 557 559 559
xxx
Contents
15.
Central Effects of General Anesthesia 571 Eckehard A.E. Stuth, Edward J. Zuperku, and Astrid G. Stucke I. II. III. IV. V. VI.
16.
The Influence of Inhalational Anesthetics on Carotid Body Mediated Ventilatory Responses Albert Dahan, Raymonda Romberg, Elise Sarton, and Luc J. Teppema I. II. III. IV. V.
17.
18.
Introduction General Effects of Anesthetics on Respiration Anesthetic Effects on Fast Synaptic Neurotransmission Overview of the Brainstem Respiratory Network Paradigms of Anesthetic Effects on Respiratory Neurotransmission Summary and Outlook Acknowledgment References
Introduction Influence of Inhalational Anesthetics on the Ventilatory Response to Hypoxia and Hypercapnia Pain and Behavioral Responses Short-Term Potentiation of Breathing (STP) Conclusions References
571 572 583 597 613 630 630 630
653
653 655 671 674 677 677
General Anesthesia and Respiratory Mechanics David O. Warner
687
I. II. III. IV. V. VI.
687 688 694 709 714 722 722
Introduction Normal Function of the Respiratory Pump Effects of Anesthesia on Chest Wall Mechanics Effects of Anesthesia on Upper Airway Mechanics Effects of Anesthesia on Lung Mechanics Summary References
Recovery from Anesthesia Shiroh Isono and Jacob Rosenberg
737
I. II.
737
III.
Introduction Impairment and Recovery of Upper Airway Function after Anesthesia Impairment and Recovery of Chemical Control of Breathing after Anesthesia
738 750
Contents IV. V.
19.
20.
xxxi Impairment and Recovery of Lung Function after Surgery Late Postoperative Nocturnal Hypoxemia References
752 755 764
Neuromuscular Blocking Agents and Ventilation Lars I. Eriksson
779
I. II.
780
Regulation of Breathing Respiratory Pump Function and the Control of the Upper Airways References
785 789
Cardiovascular Drugs and the Control of Breathing Denham S. Ward and Suzanne Karan
793
I. II. III. IV. V. VI.
793 794 800 801 802 804 805
Introduction Catecholamine Agonists and Antagonists Renin–Angiotensin System Calcium Channel Blockers Purinoceptor Agonists and Antagonists Other Agents References
Author Index Subject Index
815 823
Part I Neuropharmacology and Physiology
1 Peripheral Chemoreceptors: Sensors of Metabolic Status
COLIN A. NURSE McMaster University Hamilton, Ontario, Canada
I.
Introduction
The mammalian carotid body (CB) is a major peripheral chemoreceptor organ that helps in maintaining the chemical composition of arterial blood via the control of respiration. Consistent with its ability to sense bloodborne chemical stimuli, as originally suggested by the earlier studies of DeCastro, the organ is richly innervated and is supplied by an elaborate vascular network that has earned it the reputation as the tissue with the highest blood flow per unit weight [1,2]. Despite their small size, ranging from a few hundred microns to a few millimeters depending on species, these bilaterally paired organs occupy a strategic location near the carotid bifurcation, at the junction where the common carotid artery divides into its internal and external branches, supplying blood to the brain. Since the pioneering work of Heymans and collaborators, it is now well established that the CB is excited by a variety of chemical signals in arterial blood including low PO2, elevated PCO2 and acidic pH [2–5]. These chemoexcitants cause an increased action potential discharge in the carotid sinus nerve (CSN) whose projections to the central pattern generator in the 3
4
Nurse
brainstem ultimately lead to a compensatory increase in ventilation. There is strong evidence that the receptor cells for these stimuli are the catecholamine-producing glomus or type I cells [2,6–8], which occur in clusters and receive sensory innervation from the petrosal ganglia via the CSN [1–3]. More recent studies suggest an expanded role of the CB as a polymodal arterial chemosensor. For example, the recent demonstration that CB type I cells can respond to low glucose by secreting catecholamines [9] raises the idea that the organ might be broadly categorized as a ‘metabolic sensor’, that is, it is able to detect a range of circulating products and precursors of cellular metabolism. Moreover, these cells can also sense temperature and osmolarity [10–12] that is dependent on the concentration of circulating water and ions in plasma and extracellular fluid. These advances impinge on a wealth of information available about the transductive events in type I cells and on more recent information about the neurotransmitter mechanisms that translate the receptor potential into an increased afferent central nervous system (CNS) discharge. This review will highlight some of these recent advances. The interesting effects of chronic and intermittent stimuli (e.g., hypoxia) on CB plasticity are areas of physiological and pathophysiological significance, but will not be discussed here due to space constraints. The reader may wish to consult other recent reviews on this topic [13,14]. Also, though pulmonary neuroepithelial bodies (NEBs) represent another well-studied class of peripheral chemoreceptors that sense airway hypoxia [15,16], they have been the subject of recent reviews [17,18] and will not be considered in any detail here. II.
Cellular Organization and Innervation of the Carotid Body
The major cell types of the mammalian CB are the chemoreceptor type I cells which are organized in clusters of 20 or more cells, in intimate association with glial-like sustentacular or type II cells [1]. These chemoreceptors contain biogenic amines, especially dopamine [2,3], and accordingly express tyrosine hydroxylase (TH), the rate-limiting enzyme in catecholamine biosynthesis (Figure 1.1A,B). Other biogenic amines present in type I cells include norepinephrine and the indoleamine 5-HT or serotonin [2,19,20]. There is also substantive evidence that type I cells can synthesize and release acetylcholine in several species [2–4,21–23], though surprisingly, attempts to localize cholinergic gene expression in these cells have been unsuccessful in the rat [24]. A variety of other neurotransmitter candidates has been linked to CB chemoreceptors including substance P [5], GABA [19], and ATP [23]. The chemoreceptor clusters are surrounded by a rich network of fenestrated capillaries that permit ready access of blood-borne chemicals,
Peripheral Chemoreceptors 5
Figure 1.1 Immunofluorescence staining of a tissue section from 13-day-old rat carotid body. In A, the scattered type I cell clusters are immunopositive for tyrosine hydroxylase (TH); the TH antibody was raised in rabbit and visualized with an FITC-conjugated goat antirabbit IgG. The area enclosed by box in A, is shown in B,C after dual immunofluorescence staining for TH and neurofilament (NF), respectively. In C, a mouse anti-NF antibody was used and visualized with Texas red-conjugated goat anti-mouse IgG. The nerve processes and terminals in C (arrows) are in close apposition with the TH-positive type I cells shown in B. Calibration bars represent 40 mm in A, and 20 mm in B,C.
6
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including intravenously administered large dye molecules, e.g., Evans blue dye [25]. Gap junctions link chemoreceptor cells to each other and possibly to type II cells and sensory nerve endings [1,26], and this is likely the basis for electrical coupling observed between adjacent receptor cells [27]. Additionally, ultrastructural studies suggest chemical reciprocal synapses are present between neighboring chemoreceptor cells and between chemoreceptor cells and sensory nerve endings [1], suggesting a role for local circuits in the processing of chemosensory information in the CB. While neurofilament (NF)-positive, afferent fibers (Figure 1.1C), arising mainly from TH-positive neurons in the petrosal ganglion [28], provide the major sensory innervation to chemoreceptor cells, both autonomic and sensory fibers supply innervation to the whole organ [29]. Among these are a plexus of NO-synthesizing nerve fibers, originating from neurons within the glossopharyngeal (GPN) nerve, which are thought to provide efferent inhibition of the CB [30]. Recent evidence suggests that, in addition to causing excitation of type I cells, hypoxia can also directly excite these GPN neurons [31], thereby providing a pathway for negative feedback control of CB activity during natural stimulation.
III.
Carotid Body Chemoreceptors: O2-Sensitive Kþ Channels
Carotid body chemoreceptors or type I cells respond to hypoxia with membrane depolarization and/or increased excitability (e.g., Figure 1.2D), which facilitates entry of extracellular Ca2þ and neurosecretion [2,5–7,32,33]. Since the original description in the rabbit by Lopez-Barneo and colleagues [34], the main hypoxia-sensing mechanism in type I cells appears to involve inhibition of Kþ channels, though the particular subtype(s) of Kþ channel may differ among species and more than one O2-sensitive Kþ channel may occur in the same cell [5,8,35]. For example, inhibition of two distinct O2-sensitive Kþ channels are thought to play key roles in hypoxic chemotransduction in rat type I cells [8,36,37]. One type is the voltageand Ca2þ-dependent large conductance Kþ (BK) channel [7,8]. This is illustrated in Figure 1.2A–C, where inhibition of the BK current by removal of extracellular Ca2þ abolishes hypoxic inhibition of the voltage-dependent outward Kþ current. The other type is a voltage-insensitive background Kþ channel which shares biophysical and pharmacological properties of the acid-sensitive, 2P-domain Kþ channel, TWIK-related acid sensitive Kþ channel (TASK) [36]. While the latter alone satisfactorily accounts for membrane depolarization or receptor potential in single-isolated type I cells during hypoxia [37], both channels most likely contribute to the response of type I cells in their native clustered arrangement (Figure 1.1).
Peripheral Chemoreceptors
7
A
3000
B
2000 Control Wash Hypoxia 0 Ca2+ Hypoxia + 0 Ca2+
1 nA
1000
10 ms −120
C Hypoxia
2000
0 Ca2+ + Hypoxia
0 Ca2+
−80
−40
40
80
D
I (pA)
1600 10 mv 4s
1200 Time (s) 800
Hypoxia 0
10
20
30
Figure 1.2 Effects of hypoxia on whole-cell currents and membrane potential in rat type I cells in culture. A. Voltage step from a holding potential of 60 mV to þ 50 mV elicits an outward Kþ current which is reversibly suppressed (20%) on exposure to a hypoxic solution (PO2 ¼ 20 mmHg). In nominally Ca2þ-free solution, the outward current is suppressed by almost the same amount as hypoxia, and hypoxia had no additional effect on the residual current in Ca2þ-free solution. B. I–V relation for the cell in A shows the effects of hypoxia, Ca2þ-free solution, and both together on the whole-cell currents. C. Time-series plot of the effects of hypoxia and Ca2þ-free solution on the same cell as in A and B. These results confirm that hypoxia suppresses a Ca2þ-dependent Kþ current in rat type I cells [7,8]. D. Hypoxia evoked membrane depolarization that was sufficient to elicit action potentials in a different type I cell (Data from Ref. 33).
Evidence for this view is supported by the observations that blockade of BK channels with iberiotoxin leads to catecholamine secretion from clustered type I clusters in culture [38] and in CB tissue slices [39], suggesting BK channels are normally open under resting ‘‘normoxic’’ conditions in these clusters. Interestingly, Naþ channel density is low or absent in rat type I cells [7,40] though the hypoxia-induced depolarization or receptor potential seen in these usually quiescent cells may in some cases lead to action potentials (Figure 1.2D), presumably mediated largely by Ca2þ influx through voltage-dependent L-type Ca2þ channels [32]. However, there is evidence that spontaneous spike activity may occur in the larger clusters of rat type I cells [20], and in these cases hypoxia can cause an increase in spike
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frequency and broadening of the action potential, where the latter is thought to arise from inhibition of BK channels. In rabbit type I cells the O2-sensitive Kþ channels are different from rat, and include both a voltage-dependent, 4-AP-sensitive, transient Kþ channel [2,6,35] and an inward rectifier, HERG (human ether-a-gogorelated gene)-like Kþ channel [5]. Since rabbit type I cells show spontaneous activity, the voltage-dependent, 4-AP-sensitive Kþ channels are frequently open and their closure by hypoxia increases firing frequency and neurotransmitter release [41]. IV.
What is the O2 Sensor and How is it Linked to K1 Channel(s)?
A major unsolved issue in CB function is the molecular identity and location of the O2 sensor and the signal transduction pathway that couples the sensor to O2-sensitive Kþ channels [35]. One candidate heme protein, the neutrophil-like NADPH oxidase, which satisfies the main criteria for the O2 sensor in pulmonary neuroepithelial bodies [16], does not play this role in type I chemoreceptor cells [42,43, our unpublished observations]. Other candidates include heme proteins in the mitochondrial electron transport chain and heme proteins that are closely associated with the O2-sensitive plasma membrane Kþ channels [35,41,44]. In cell-attached patches of rat type I cells, hypoxia closes TASK-1like Kþ channels, but regulation is lost in excised inside-out patches, suggesting a cytoplasmic messenger or cofactor is required for conferring O2-sensitivity [36]. A suggestion that mitochondria might be involved in the link between the O2 sensor and plasma membrane TASK-1-like Kþ channels in these cells was indicated by the observation that inhibitors of the electron transport chain mimic the effects of hypoxia on background Kþ channels [45]. On the other hand, in rabbit type I cells, hypoxic modulation of single Kþ channels has been observed in excised membrane patches [46], suggesting that these channels may either possess intrinsic O2-sensitivity, or may be closely associated with a separate O2-sensor, e.g., hemoprotein. Thus, there remains the question whether there is a single O2-sensor in type I cells regardless of the Kþ channel type being modulated, or whether there may be more than one sensor, even in the same cell, for different O2-sensitive Kþ channels. V.
Role of Fast-Acting Neurotransmitters in Chemosensory Processing
While strong evidence that CB type I cells act as chemosensors for hypoxic, hypercapnic, and acidic stimuli has accumulated over the last roughly
Peripheral Chemoreceptors
9
15 years [2,7,8], there has been slower progress on the neurotransmitter mechanisms that translate the receptor potential in type I clusters to an increased afferent discharge in the CSN. Dopamine (DA), one of the best studied CB neurotransmitters, had received much attention as a leading transmitter candidate in chemoreception [2], but fell into disfavor with the demonstration that depletion of CB dopamine had little or no effect on the hypoxia-induced increase in CSN discharge [47,48]. The more popular view to date is that DA acting on presynaptic and possibly postsynaptic D2 dopamine receptors likely plays an inhibitory role in modulating CB function [24]. On the other hand, acetylcholine (ACh) has for many years remained an attractive candidate for mediating CB chemoexcitation [3,4]. However, the skeptics were not satisfied because blockers of both nicotinic and muscarinic ACh receptors could not inhibit completely the CB response to natural stimulation [4]. A satisfactory solution to this impasse was recently provided with the demonstration that co-release of ATP and ACh was likely the main mechanism mediating hypoxic chemotransmission in the rat CB [23]. These studies were greatly facilitated by the development of a co-culture model in which rat type I cell clusters formed de novo functional synaptic connections with dissociated petrosal (afferent) neurons in vitro [21,49]. The main advantage of this co-culture preparation, compared with the commonly used isolated CB-sinus nerve preparation, was that subthreshold postsynaptic responses could be recorded from neuronal cell bodies that were fortuitously juxtaposed to type I clusters [21–23,49,50]. As illustrated in Figure 1.3A,B, transmission of two natural CB stimuli, i.e., hypoxia and isohydric hypercapnia, can be demonstrated in these co-cultures and in both cases the postsynaptic response recorded in the neuron was reversibly inhibited by reduction of the extracellular Ca2þ: Mg2þ ratio. These results are consistent with the idea that chemical synaptic transmission is indeed required and that the responses are not due to a direct action of the chemostimuli on the neurons. In Figure 1.3A, the neuronal soma was close enough to the type I cluster to allow recording of the postsynaptic depolarization, which in this case was near the spike threshold and caused a burst of action potentials. In this configuration, soma recordings provide a replica of synaptic events occurring at the nerve terminals. The equivalent experiment in situ would require electrophysiological recordings from intact nerve terminals apposed to CB type I cells. This experimental approach has only met with limited success due to technical difficulties in obtaining stable recordings from these nerve endings in situ [51]. The ability to record the postsynaptic depolarization following application of a chemosensory stimulus in co-culture permitted pharmacological identification of the neurotransmitters involved. As illustrated
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A 20 mV 2s
Hypoxia
Hypoxia
Hypoxia
0.1 mM Ca, 6 mM Mg
20 mV 4s
B
10% CO2
10% CO2
10% CO2
Figure 1.3 Role of chemical transmission in mediating the effects of CB chemostimuli on petrosal neurons that functionally innervated type I clusters in co-culture. Perforated patch recordings of membrane potential were obtained from petrosal neurons that were adjacent to a type I cluster. In A, hypoxia (PO2 5 mmHg; duration indicated by horizontal bar) caused depolarization and a burst of action potentials in a functional petrosal neuron and the effect was reversibly abolished when the extracellular Ca:Mg ratio was decreased from 2 mM Ca2þ:1 mM Mg2þ (left and right traces) to 0.1 mM Ca2þ:6 mM Mg2þ (middle trace). A similar result was obtained when isohydric hypercapnia (10% CO2; pH ¼ 7.4) was used as the chemosensory stimulus in a different co-culture (B). Note spontaneous activity in the co-cultured neuron in B, under control (normocapnic) conditions (5% CO2; pH ¼ 7.4), i.e., to the left and right of the horizontal bars. These data suggest that chemical transmission is involved in the transfer of chemosensory information from the type I cluster to the neurons. The neuronal resting potential was 55 mV in A and B.
in Figure 1.4A, the hypoxia-induced postsynaptic response could be completely inhibited by the combined presence of a nicotinic ACh receptor blocker (100 mM hexamethonium) and a purinergic receptor blocker (50 mM suramin). Similar results were obtained when isohydric and acidic hypercapnia were used as chemostimuli in functional co-cultures (50, our unpublished observations). Other nicotinic and purinergic blockers successfully used in these co-cultures include mecamylamine (1–2 mM) and reactive blue 2 (10–50 mM), respectively [22,49]. In almost all cases, even high concentrations of a single blocker were insufficient to abolish
Peripheral Chemoreceptors
11
A 20 mV 50 µM hex + 25 µM sur
Hypoxia
B
100 µM hex + 50 µM sur
Hypoxia
Hypoxia
2s
Hypoxia
50 µM hex + 25 µM sur
Hypoxia
Hypoxia
20 s
Hypoxia
Figure 1.4 Evidence that co-release of ACh and ATP mediates hypoxic chemotransmission in co-cultured petrosal neurons, and in the isolated carotid body-sinus nerve preparation in the rat. In A, combined application of hexamethonium (hex) and suramin (sur), blockers of nicotinic ACh and purinergic receptors respectively, caused a dose-dependent inhibition of postsynaptic excitatory response recorded in a petrosal neuron, juxtaposed to a type I cluster (Data from Ref. 23). Similarly, in the intact CB–sinus nerve preparation in vitro (B), the hypoxiainduced sensory discharge recorded extracellularly in the sinus nerve was reversibly abolished in the combined presence of hexamethonium and suramin. The effect of either drug alone produced only partial inhibition of the sensory discharge (not shown). These data support the idea that co-release of ACh and ATP is the major mechanism that mediates hypoxic signaling in the rat carotid body.
the hypoxic response though partial inhibition was consistently obtained [23,49]. The conclusion that co-release of ACh and ATP from type I cells was the principal mechanism underlying hypoxic chemotransmission was not biased by the artificial culture conditions, since in the intact rat CB– sinus nerve preparation the same combination of nicotinic and purinergic blockers was required to abolish the chemosensory discharge, recorded extracellularly in the sinus nerve (Figure 1.4B; [23]). While the molecular composition of the functional postsynaptic AChR must yet be determined, there is strong evidence based on electrophysiological, molecular (RT-PCR), and immunocytochemical techniques that the functional purinergic receptor is a heteromultimer of P2X2–P2X3 subunits [23,50]. Both nicotinic and purinergic receptors are co-expressed in chemosensory neurons where they function as fast-acting, ligand-gated ion channels [23]. Importantly, immunoreactive P2X2 and P2X3 subunits co-localize in sensory nerve terminals apposed to type I receptor cells in the rat CB in situ [23].
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Glucose Sensing in the Carotid Body
It was recently demonstrated that low glucose stimulates catecholamine (CA) release from type I cells in tissue slices of the rat CB [9]. Similar to other CB chemostimuli, the effect of low glucose on CA secretion was dosedependent and was abolished by blockers of voltage-gated Ca2þ channels, suggesting that the response depended on membrane depolarization [9]. While low glucose selectively inhibited voltage-dependent outward Kþ current in rat type I cells, the fact that the effect persisted after blockade of the O2-sensitive Ca2þ-dependent maxi-Kþ (BK) current with iberiotoxin indicated a transduction pathway distinct from the one used to signal hypoxia [9]. These data suggest that type I cells are also physiological glucose sensors and, as such, are capable of integrating sensory information arising from a variety of chemical signals that contribute to general metabolic status. Integration of these signals would be expected to influence the frequency of afferent sinus nerve discharge and subsequent activation of the cardiorespiratory and cardiovascular system. Since in our co-culture experiments discussed above, recordings from functional petrosal neurons, co-cultured with type I receptor clusters, were routinely carried out in the presence of high (10 mM) glucose [23], we decided to test whether or not glucose withdrawal (substituted with equimolar sucrose) could stimulate neuronal activity. Indeed, as illustrated in Figure 1.5, the same chemosensory unit that responded to hypoxia with a reversible increase in neuronal spike discharge was similarly excited following glucose withdrawal. Moreover, this postsynaptic excitatory response induced by low glucose was inhibited by combined application of mecamylamine and suramin (our unpublished observations), suggesting that, as was the case for hypoxia (see above), co-release of ACh and ATP from type I cells was required for signaling low glucose. Though other organs are involved in the sensing of blood glucose [52], the strategic location of the CB may be important for brain function since neurons are especially sensitive to simultaneous glucose and O2 deficiency [9]. VII.
Neuromodulation in the Carotid Body
A contributing factor to the slow progress made in identifying the key neurotransmitters that mediate CB chemoexcitation is the wealth of transmitter candidates localized to type I cells [2]. There is evidence that several of them play modulatory roles, where the effect may be either presynaptic, i.e., at the level of the type I cell, or postsynaptic, i.e., at the sensory nerve endings. For example, one presynaptic effect of dopamine acting via D2 receptors on type I cells is to inhibit L-type Ca2þ channels [53]. Such an effect could lead to a negative feedback regulation of
Peripheral Chemoreceptors Control
13 Hypoxia
0 glucose
Wash
20 mV 2s
Figure 1.5 Chemosensory units in co-culture are excited by both hypoxia and low extracellular glucose. Recordings were obtained from a petrosal neuron that was near a type I cluster in co-culture. Note that the neuron, which was quiescent under control normoxic conditions (upper left and right traces), increased firing when exposed to hypoxia (upper middle trace). In this same neuron, removal of extracellular glucose (10 mM glucose that is normally present in the recording medium was replaced by 10 mM sucrose) also caused excitation in the neuron (lower middle trace) and the effect was reversible. Thus, the same chemosensory unit can be excited by hypoxia and low glucose.
neurotransmitter release during hypoxia by an autocrine–paracrine mechanism. Our recent studies suggest that 5-HT (serotonin) and GABA, which are both expressed in rodent type I cells [19,20,54,55], may act as positive and negative modulators of type I chemoreceptor function, respectively. We found that the 5-HT receptor blocker, ketanserin, inhibited the spontaneous spike discharge that is occasionally seen in large type I clusters in culture, and exogenous 5-HT can induce membrane depolarization or rhythmic-like spiking in type I clusters [20]. The effect of 5-HT appears to be mediated via G-protein-coupled 5-HT2a autoreceptors and protein kinase C-dependent inhibition of Kþ channels [55]. Whereas 5-HT can potentiate transmitter output from type I clusters by autocrine–paracrine mechanisms, GABA has the opposite effect. In particular, GABA appears to inhibit transmitter output via G-protein coupled GABAB autoreceptors and PKAdependent augmentation of a TASK-1-like background Kþ current [54]. Other neuromodulators are likely to play important roles in the regulation of type I cell function. For example, ACh activates both nicotinic and muscarinic autoreceptors which are known to be expressed in type I cells of several species [4]. There is also the possibility that ATP, which as described above is an important fast-acting transmitter that is co-released with ACh during CB chemoexcitation, may act presynaptically as well. So far, we have been unable to detect ATP sensitivity in type I cells in
14
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Type I
100 pA 20 ms
5 mV 4s
50 µM ATP
50 µM ATP
Figure 1.6 Comparison of ATP sensitivity in cultured rat CB type I and type II cells. Top traces show voltage clamp recordings of ionic currents in a type II cell (left) and a type I cell (right) after a few days in culture. Note the absence of voltagedependent outward Kþ currents in the type II cell, but their presence in the type I cell. Currents were evoked during 10 mV incremental voltage steps from 180 mV to þ 60 mV; holding potential ¼ 60 mV. Lower traces show current clamp recordings of membrane potential in the same type II and type I cells (as in upper traces) after rapid perfusion of ATP over the soma. Note that ATP causes membrane depolarization in the type II, but not type I cell. The resting potential was 40 mV in the type II cell and 47 mV in the type I cell.
voltage or current clamp studies (e.g., Figure 1.6). However, the possibility that ATP may influence intracellular Ca2þ homeostasis in type I cells via regulation of intracellular Ca2þ stores is not excluded. Interestingly, we have preliminary data indicating that sustentacular or type II cells, which are intimately associated with type I cell clusters in situ [1] and in culture [56], may express ATP receptors (Figure 1.6). Further studies are required to determine the molecular identity of these purinergic receptors, though the possibility is raised that type II cells may contribute to the overall autocrine regulation of CB function. For example, type II cells may help spread electrical activity within type I clusters via their long, slender processes which tend to surround and penetrate the clusters [56]. ATP released during chemosensory transmission
Peripheral Chemoreceptors
15
may also contribute indirectly to autocrine–paracrine signaling in the CB after its enzymatic breakdown to adenosine by ectonucleotidases. Adenosine, derived in this way or as a circulating metabolite whose levels in plasma increase during hypoxia, may itself act on adenosine A2A autoreceptors on type I cells and contribute to CB excitation [57], perhaps via inhibition of a 4-AP-sensitive, voltage-dependent outward Kþ current [58]. Thus, the purine ATP may be a central player in CB function, acting directly as a fast co-transmitter in mediating the postsynaptic response and as a presynaptic neuromodulator via type II cells, or indirectly, as a modulator of type I cell function via its breakdown product, adenosine. VIII.
Future Directions
One of the key issues that needs to be clarified is the molecular identity and location of the O2 sensor in CB chemoreceptors and the signaling pathway leading to Kþ current inhibition in low oxygen. Current methodologies using patch clamp techniques at the whole-cell and single-channel level, carbon fiber amperometry to assay for amine secretion in tissue slices, and the application of mitochondrial inhibitors will need to be complemented by functional genomic approaches where the sensor function can be directly manipulated. It will also be of interest to identify the glucose sensor in these cells and the transductive process leading to increased secretion from type I cells in low extracellular glucose. The complexity surrounding the need for such a broad range of neurotransmitters/neuromodulators in type I cells is still puzzling; however, the possibility that different chemostimuli might release a different array of these chemical signals needs further exploration. Finally, since these neurotransmitters and their receptors appear to endow the organ with a remarkable ability to fine tune its responses to any particular stimulus, their likely contribution to chemoreceptor plasticity following chronic stimulation needs to be explored in the future. Acknowledgments The work attributed to my laboratory was supported by grants from the Canadian Institutes of Health Research. I wish to thank Cathy Vollmer for expert technical assistance, Min Zhang for carrying out many of the experiments on co-cultures and providing most of the figures illustrated in the text, Huijun Zhong for providing the data for Figure 1.2, and Mike Jonz for assistance with Figure 1.1. I am indebted to several colleagues for contributing to the ideas expressed in this article, particularly Min Zhang, Veronica Campanucci, and Ian Fearon.
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Nurse References
1. 2.
3. 4.
5. 6. 7. 8. 9. 10. 11. 12.
13. 14.
15. 16.
17. 18.
19.
McDonald, D.M., Peripheral chemoreceptors, in Regulation of Breathing, Part I, Hornbein, T.F., ed., New York, Marcel Dekker, Inc., pp. 105–319, 1981. Gonzalez, C., Almaraz, L., Obeso, A. and Rigual, R., Carotid body chemoreceptors: from nature stimuli to sensory discharges, Physiol. Rev. 74, 829–898, 1994. Eyzaguirre, C. and Zapata, P., Perspectives in carotid body research, J. Appl. Physiol. 57, 931–957, 1984. Fitzgerald, R.S., Oxygen and carotid body chemotransduction: the cholinergic hypothesis—a brief history and new evaluation, Resp. Physiol. 120, 89–104, 2000. Prabhakar, N.R., Oxygen sensing by the carotid body chemoreceptors, J. Appl. Physiol. 88, 2287–2295, 2000. Lopez-Barneo, J., Oxygen-sensing by ion channels and the regulation of cellular functions, Trends Neurosci. 19, 435–440, 1996. Peers, C. and Buckler, K.J., Transduction of chemostimuli by the type I carotid body cell, J. Memb. Biol. 144, 1–9, 1995. Peers, C., Oxygen-sensitive ion channels, Trends Pharmacol. Sci. 18, 405–408, 1997. Pardal, R. and Lopez-Barneo, J., Low glucose-sensing cells in the carotid body, Nature Neurosci. 5, 197–198, 2002. Gallego, R., Eyzaguirre, C. and Monti-Bloch, L., Thermal and osmotic responses of arterial receptors, J. Neurophysiol. 42, 655–680, 1979. Carpenter, E. and Peers, C., Swelling- and cAMP-activated Cl currents in isolated rat carotid body type I cells, J. Physiol. 503, 497–511, 1997. Molnar, Z., Petheo, G.L., Fulop, C. and Spat, A., Effects of osmotic changes on the chemoreceptor cell of rat carotid body, J. Physiol. 546, 471– 481, 2003. Prabhakar, N.R., Oxygen sensing during intermittent hypoxia: cellular and molecular mechanisms, J. Appl. Physiol. 90, 1986–1994, 2001. Wang, Z.-Y. and Bisgard, G.E., Chronic hypoxia-induced morphological and neurochemical changes in the carotid body, Micros. Res. Tech. 59, 168–177, 2002. Youngson, C., Nurse, C., Yeger, H. and Cutz, E., Oxygen sensing in airway chemoreceptors, Nature 365, 153–155, 1993. Fu, X.W., Wang, D., Nurse, C., Dinauer, M.C. and Cutz, E., NADPH oxidase is an O2 sensor in airway chemoreceptors: evidence from Kþ current modulation in wild type and oxidase-deficient mice, Proc. Natl. Acad. Sci. USA 97, 4374–4379, 2000. Cutz, E. and Jackson, A., Neuroepithelial bodies as airway oxygen sensors, Respir. Physiol. 115, 201–214, 1999. Kemp, P.J., Lewis, A., Hartness, M.E., Searle, G.J., Miller, P., O’Kelly, I. and Peers, C., Airway chemotransduction: from oxygen sensor to cellular effector, Am. J. Respir. Crit. Care Med. 166, 517–524, 2002. Oomori, Y., Nakaya, K., Tanaka, H., Iuchi, H., Ishikawa, K., Satoh, Y. and Ono, K., Immunohistochemical and histochemical evidence for the presence of
Peripheral Chemoreceptors
20.
21.
22.
23.
24. 25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
17
noradrenaline, serotonin and gamma-aminobutyric acid in chief cells of the mouse carotid body, Cell Tiss. Res. 278, 249–254, 1994. Zhang, H. and Nurse, C.A., Does endogenous 5-HT mediate spontaneous rhythmic activity in chemoreceptor clusters of rat carotid body? Brain Res. 872, 199–203, 2000. Zhong, H., Zhang, M. and Nurse, C.A., Synapse formation and hypoxic signalling in co-cultures of rat petrosal neurones and carotid body type 1 cells, J. Physiol. 503, 599–612, 1997. Nurse, C.A. and Zhang, M., Acetylcholine contributes to hypoxic chemotransmission in co-cultures of rat type 1 cells and petrosal neurones, Respir. Physiol. 115, 189–199, 1999. Zhang, M., Zhong, H., Vollmer, C. and Nurse, C.A., Co-release of ATP and ACh mediates hypoxic signalling at rat carotid body chemoreceptors, J. Physiol. 525, 143–158, 2000. Gauda, E.B., Gene expression in peripheral arterial chemoreceptors, Micros. Res. Tech. 59, 153–167, 2002. McDonald, D.M. and Blewett, R.W., Location and size of carotid-body like organs (paraganglia) revealed in rats by permeability of blood vessels to Evans blue dye, J. Neurocytol. 10, 607–643, 1981. Kondo, H., Are there gap junctions between chief (glomus, type I) cells in the carotid body chemoreceptors? A review, Micros. Res. Tech. 59, 227–233, 2002. Abudara, V. and Eyzaguirre, C., Electrical coupling between cultured glomus cells of the rat carotid body: observations with current and voltage clamping, Micros. Res. Tech. 59, 249–255, 2002. Katz, D.M. and Black, I.B., Expression and regulation of catecholaminergic traits in primary sensory neurones: relationship to target innervation in vivo, J. Neurosci. 6, 983–989, 1986. Ichikawa, H., Innervation of the carotid body: immunohistochemical, denervation, and retrograde tracing studies, Micros. Res. Tech. 59, 188–195, 2002. Wang, Z.Z., Stensaas, L.J., Dinger, B.G. and Fidone, S.J., Nitric oxide mediates chemoreceptor inhibition in the cat carotid body, Neuroscience 65, 217–229, 1995. Campanucci, V.A., Fearon, I.M. and Nurse, C.A., A novel O2-sensing mechanism in glossopharyngeal neurones mediated by a halothane-inhibitable background Kþ conductance, J. Physiol. 548, 731–743, 2003. Buckler, K.J. and Vaughan-Jones, R.D., Effects of hypoxia on membrane potential and intracellular calcium in rat neonatal carotid body type I cells, J. Physiol. 476, 423–428, 1994. Zhong, H., Electrophysiology and Transmitter Sensitivities of Isolated Rat Petrosal Neurons: Synapse Formation and Hypoxic Signaling in Co-culture with Carotid Body Chemoreceptors, Ph.D. Dissertation, McMaster University, Hamilton, Ontario, 1997. Lopez-Barneo, J., Lopez-Lopez, J.R., Urena, J. and Gonzalez, C., Chemotransduction in the carotid body: Kþ current modulated by PO2 in type I chemoreceptor cells, Science 241, 580–582, 1988.
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Nurse
35.
Lopez-Barneo, J., Pardal, R. and Ortega-Saenz, P., Cellular mechanisms of oxygen sensing, Ann. Rev. Physiol. 63, 259–287, 2001. Buckler, K.J., Williams, B.A. and Honore, E., An oxygen-, acid- and anaesthetic-sensitive TASK-like background potassium current in rat arterial chemoreceptor cells, J. Physiol. 525, 135–142, 2000. Buckler, K.J., A novel oxygen-sensitive potassium current in rat carotid body type I cells, J. Physiol. 498, 649–662, 1997. Jackson, A. and Nurse, C., Dopaminergic properties of cultured rat carotid body chemoreceptors grown in normoxic and hypoxic environments, J. Neurochem. 69, 645–654, 1997. Pardal, R., Ludewig, U., Garcia-Hirschfeld, J. and Lopez-Barneo, J., Secretory responses of intact glomus cells in thin slices of rat carotid body to hypoxia and tetraethylammonium, Proc. Natl. Acad. Sci. USA, 97, 2361–2366, 2000. Stea, A. and Nurse, C.A., Whole-cell and perforated-patch recordings from O2-sensitive rat carotid body cells grown in short- and long-term culture, Pflu¨gers Archiv. 418, 93–101, 1991. Lopez-Barneo, J., Ortega-Saenz, P., Molina, A., Franco-Obregon, A., Urena, J. and Castellano, A., Oxygen sensing by ion channels, Kidney Int. 51, 454–461, 1997. Roy, A., Rozanov, C., Mokashi, A., Daudu, P., Al-mehdi, A.B., Shams, H. and Lahiri, S., Mice lacking in gp91 phox subunit of NAD(P)H oxidase showed glomus cell [Ca2þ]i and respiratory responses to hypoxia, Brain Res. 872, 188–193, 2000. He, L., Chen, J., Dinger, B., Sanders, K., Sundar, K., Hoidal, J. and Fidone, S., Characteristics of carotid body chemosensitivity in NADPH oxidase-deficient mice, Am. J. Physiol. Cell Physiol. 282, C27–C33, 2002. Kummer, W. and Yamamoto, Y., Cellular distribution of oxygen sensor candidates—oxidases, cytochromes, Kþ-channels—in the carotid body, Micros. Res. Tech. 59, 234–242, 2002. Buckler, K.J. and Vaughan-Jones, R.D., Effects of mitochondrial uncouplers on intracellular calcium, pH and membrane potential in rat carotid body type I cells, J. Physiol. 513, 819–833, 1998. Ganfornina, M.D. and Lopez-Barneo, J., Single Kþ channels in membrane patches of arterial chemoreceptor cells are modulated by O2 tension, Proc. Natl. Acad. Sci. USA 88, 2927–2930, 1991. Donnelly, D.F., Chemoreceptor nerve excitation may not be proportional to catecholamine secretion, J. Appl. Physiol. 81, 657–664, 1996. Iturriaga, R., Alcayaga, J. and Zapata, P., Dissociation of hypoxia-induced chemosensory responses and catecholamine efflux in cat carotid body superfused in vitro, J. Physiol. 497, 551–564, 1996. Nurse, C.A. and Zhang, M., Synaptic mechanisms during re-innervation of rat arterial chemoreceptors in co-culture, Comp. Biochem. Physiol. Part A, 130, 241–251, 2001. Prasad, M., Fearon, I.M., Zhang, M., Laing, M., Vollmer, C. and Nurse, C.A., Expression of P2X2 and P2X3 receptor subunits in rat carotid body afferent neurones: role in chemosensory signalling, J. Physiol. 537, 667–677, 2001.
36.
37. 38.
39.
40.
41.
42.
43.
44.
45.
46.
47. 48.
49.
50.
Peripheral Chemoreceptors 51. 52. 53.
54.
55.
56. 57. 58.
19
Hayashida, Y. and Hirakawa, H., Electrical properties of chemoreceptor elements in the carotid body, Micros. Res. Tech. 59, 243–248, 2002. Thorens, B., A gene knockout approach in mice to identify glucose sensors controlling glucose homeostasis, Pflu¨gers Archiv. 445, 482–490, 2003. Benot, A. and Lopez-Barneo, J., Feedback inhibition of Ca2þ currents by dopamine in glomus cells of the carotid body, Eur. J. Neurosci. 2, 809–812, 1990. Fearon, I.M., Zhang, M., Vollmer, C. and Nurse, C.A., GABA mediates autoreceptor feedback inhibition in the rat carotid body via presynaptic GABAB receptors and TASK-1, J. Physiol. 553, 83–94, 2003. Zhang, M., Fearon, I.M., Zhong, H. and Nurse, C.A., Presynaptic modulation of rat arterial chemoreceptor function by 5-HT: role of Kþ channel inhibition via protein kinase C, J. Physiol. 551, 825–842, 2003. Nurse, C.A. and Fearon, I.M., Carotid body chemoreceptors in dissociated cell culture, Micros. Res. Tech. 59, 249–255, 2002. Sebastiao, A.M. and Ribeiro, J.A., Adenosine A2 receptor-mediated excitatory actions in the nervous system, Prog. Neurobiol. 48, 167–189, 1996. Vandier, C., Conway, A.F., Landauer, R.C. and Kumar, P., Presynaptic action of adenosine on a 4-aminopyridine-sensitive current in the rat carotid body, J. Physiol. 515, 419–429, 1999.
2 Central Chemoreceptors
LUC J. TEPPEMA and ALBERT DAHAN Leiden University Medical Center Leiden, The Netherlands
I.
Introduction
Some 50 years ago, Leusen, in his ventriculocisternal perfusion experiments in dogs, showed that variations in hydrogen ion concentration in the cerebrospinal fluid (CSF) had a pronounced influence on pulmonary ventilation [1]. These pioneer experiments were followed by the studies of Pappenheimer and co-workers in goats in which they showed that ventilation could be expressed as a unique function of a calculated extracellular pH existing at a location about three-fourths along the artificial steady-state concentration gradient of bicarbonate between the CSF and brain capillary plasma [2,3]. Following these classical studies, this view of central CO2 chemoreceptors that are uniquely sensitive to the pH in their microenvironment was also used by other groups in their attempts to define the ‘functional’ or ‘anatomical’ location of the central chemoreceptors (i.e., their relative distance between CSF and blood and their distance from the irregularly folded ventral surface of the medulla oblongata, respectively, e.g., [4–6]).
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Teppema and Dahan
V VI
VII VIII M IX X XI
S
L
XII
C1
Figure 2.1 Ventral medullary surface in the cat with the three ‘‘classical’’ chemosensitive areas: M (Mitchell’s area), S (Schlaefke’s area) and L (Loeschcke’s area).
A major breakthrough in the localization of the chemosensitive regions was achieved by Mitchell, Loeschcke and Schlaefke and co-workers who described three separate superficial (at a depth between 0 and 400 mm) areas on the ventral medullary surface of the cat not only containing pHsensitive cells but also neurons that were crucial in the integration of the afferent central chemosensory input to the respiratory centers (Figure 2.1, [7,8]). For many investigators these studies were the basis to further explore the ventral medullary surface of several species both in vitro and in vivo using local lesion, cooling, electrophysiological means and pharmacological agents administered by way of perfusion, superfusion, or topical application. Meanwhile, from several studies in intact animals it became clear that the assumed role of an extracellular pH as the sole stimulus was no longer tenable [9–11]. Despite this, the existence of chemosensitive regions in several species (e.g., rat, goat) analogous to those described in the cat was never seriously disputed. However, even within these chemosensitive areas, it was very difficult to identify individual respiratory CO2 chemoreceptors defined as (non-respiratory?) sensory neurons that are activated in vivo during hypercapnia (or inhibited during hypocapnia), respond directly
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(i.e., not transsynaptically) to CO2/Hþ and convey their afferent input to the respiratory centers. To date, a universal picture of respiratory CO2/Hþ chemoreceptors with characteristic morphological, neurochemical and electrophysiological properties has not yet been composed. This may have several reasons. One or more of the above criteria may be false or too loose and not specific enough. For example, in vivo and in vitro data indicate that respiratory neurons themselves may behave as central chemoreceptors [12,13], suggesting that not all chemoreceptors are purely sensory neurons. In addition, some chemoreceptors may be inhibited rather than stimulated by a decrease in pH as has been demonstrated for neurons in the ventromedial medulla [14,15]. It is also unknown if central chemoreceptors display a respiratory rhythm: some may and some may not, and they may be a heterogeneous population of cells consisting of both respiratory and nonrespiratory neurons. Another complicating factor is that there are many ways to depolarize or stimulate central chemoreceptors, but a unique intracellular factor on which all these stimuli converge and that we could use as the most direct tool to localize them, has not yet been isolated. Furthermore, the central chemoreceptors are intermingled with other brainstem neurons, complicating their identification. Finally, most of the techniques that were used may have too low a resolution to identify chemoreceptors at the singlecell level. Over the last decade, new techniques have been designed and, due to the enormous progress of molecular biology, it is now possible to use alternative approaches and to study brain function with much higher resolution. This has led to a much broader understanding of the central chemoreceptors and has shed a new light on their distribution: widespread within the brain stem and even more rostral vs. the classical view of chemosensitive regions limited to three areas on the ventral medullary surface. It also becomes progressively more evident that not only the intracellular pH may play a key role in the chemosensitive mechanism, but also ion-specific membrane channels, particularly potassium and possibly also calcium channels. By using the appropriate techniques and experimental paradigms, it is now within our reach to achieve a better understanding of the physiological role of the central chemoreceptors in health and disease. In this chapter it is our intention to focus on some major tools used over the last decade to locate or identify the central chemoreceptors and to summarize the state of the art in this regard. Secondly, we will discuss some recent data that may shed new light on the mechanism of CO2 chemoreception. Finally, we will briefly discuss the physiological role of the central chemoreceptors in the control of breathing.
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Teppema and Dahan II.
Location of Central Chemoreceptors
A. Functional and Anatomical Location
Initially, in the search for the central chemoreceptors the terms functional and anatomical location were used to define their relative distance between CSF and blood and their distance from the irregularly folded ventral surface of the medulla oblongata, respectively (see above). Both definitions are vague and may give rise to confusion. Replacing functional location by relative position or distance to blood vessels is also not very practical. Because some central chemoreceptors are more closely apposed to vessels than others, it is impossible to determine a location of chemoreceptors relative to vessels that is representative for all members. Assuming that one common qualitative feature of all chemoreceptors refers to their stimulus environment, it might be more useful to discuss whether the chemoreceptors measure a parameter that represents the PCO2 of arterial blood or rather one that reflects the tissue PCO2. In the next section we will focus on this aspect. A more practical means to estimate the distance from the medullary surface would be to describe the exact topographical anatomical location of the chemoreceptors without overlooking the sometimes considerable distance between their somata and chemosensitive (dendritic) sites. We will end this section by giving a brief overview of some major techniques used over the last decade to find the anatomical regions that contain chemoreceptors and to localize individual chemosensitive neurons. B. Central Chemoreceptors Measure Tissue PCO2
Studies in rat have shown that putative chemoreceptor cells, particularly in superficial ventral medullary tissue, are very closely associated with blood vessels. Okada et al. [16] described an intimate anatomic relationship between cells that were activated by hypercapnia (positive staining for Fos, even after synaptic blockade; see below) and surface vessels. Confirming and extending previous observations of Gorcs et al. [17], Bradley and co-workers [18], using confocal imaging and electron microscopy, showed a close association between processes of medullary serotonergic raphe neurons and large arteries in areas almost devoid of veins. Because this close association between serotonergic neurons and arteries occurs both in medullary and pontine raphe nuclei and in mesencephalon, and because many of these neurons are very sensitive to changes in pH, the authors suggested that they function as central chemoreceptors involved in ventilatory control to maintain pH/PCO2 homeostasis. The close proximity of these serotonergic cells and their
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processes to brainstem arteries in regions nearly devoid of veins would provide an optimal means to monitor a local CO2 concentration that is relatively unaffected by metabolism and would thus be an ideal location for monitoring the effectiveness of lung ventilation [18]. It is quite possible that the chemosensitive cells described by Bradley et al. function as central respiratory chemoreceptors, but not because they measure arterial PCO2 (this is already provided by the arterial chemoreceptors with their strategic location at the port of the brain circulation). Because the arterial PCO2 has no unique relationship with alveolar ventilation (this relationship alters with changes in metabolism and inspired CO2), it cannot be considered a unique parameter that represents the level or effectiveness of alveolar ventilation. Monitoring a variable that reflects the tissue PCO2 would provide more useful information. According to the mass balance for CO 2 , the P CO 2 in brainstem tissue (where the chemoreceptors are located) is determined by local metabolism and blood flow; CO2 reactivity of brainstem vessels; the arterial PCO2; the slopes of the CO2 dissociation curves for blood and tissue, and a parameter
that locates the chemoreceptor PCO2 somewhere between the tissue and arterial PCO2 [19–21]. For this very reason, by measuring the tissue PCO2, the central chemoreceptors provide an optimal feedback signal for metabolic ventilatory control. Thus, chemoreceptors that measure a parameter that closely reflect arterial CO2 concentration before it has been influenced by local tissue metabolism [18] will not provide the most relevant feedback signal for the metabolic control of breathing. The remaining question now is: do the putative respiratory chemoreceptors described by Bradley et al. [18] measure an arterial or tissue PCO2? Assuming that the processes of these chemosensitive cells measure a PCO2/[Hþ] in their immediate environment, i.e., the perivascular tissue space (with a relatively low value of parameter ), then these cells will measure a variable that must also reflect local metabolism simply because (in the case of hypercapnia) the amount of CO2 that diffuses out from local blood vessels will depend on local blood flow and on the amount of metabolically produced CO2 already present. That indeed the tissue PCO2 is the measured variable rather than a parameter closely representing the arterial PCO2 has been shown in numerous studies in men and whole animals. We give a small selection of numerous examples: 1.
In an early study in anesthetized dogs, Dutton et al. [22] found that following perfusion of the vertebral arteries with hypercapnic blood for 2 minutes, the rate of recovery of ventilation was much faster than after steady-state inhalation of CO2, a finding indicating that the central chemoreceptor PCO2 is dominated by the tissue and CSF PCO2 rather than by the PCO2 in capillary blood. Studies using the technique of dynamic end-
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Teppema and Dahan tidal forcing showed that the time course of the central chemoreflex loop to step-changes in end-tidal PCO2 or arterial PCO2 of blood selectively perfusing the brainstem does not reflect that of the (near) step in arterial PCO2 but rather is determined by the speed of CO2 loading and unloading of the brainstem [23–25]. 2. In cats undergoing artificial perfusion of their brainstems, a selective increase in medullary blood flow induced by adding the vasodilator papaverine to the central blood at constant arterial peripheral and central blood gas tensions, resulted in a decrease in ventilation due to increased washout of CO2 from the pontomedullary region [26]. In this preparation at constant peripheral blood gas tensions, when the PO2 of the blood perfusing the brainstem was lowered from a hyperoxic to a normoxic level without changing its PCO2, ventilation decreased with a time constant determined by the increased washout of CO2 [27]. 3. In the cat, the carbonic anhydrase inhibitors benzolamide and acetazolamide, administered systemically in a dose sufficiently large to completely inhibit erythrocytic carbonic anhydrase, a decrease in the end-tidal and arterial PCO2 in vivo is followed (not preceded) by a large rise in ventilation that is accompanied by a considerable brain stem tissue acidosis (rise in PCO2 and fall in pH) in the face of a lower arterial PCO2 of the blood perfusing the central chemoreceptors [28–30].
If the measured parameter is a reflection of the tissue PCO2, does it mean that all chemoreceptors measure an equal P CO 2 or pH? The answer will be no, because one of the determinants of local PCO2 is local blood flow which will not be equal in all chemosensitive regions. This is illustrated by the existence of appreciable pH gradients in the medulla oblongata as demonstrated in the cat [31]. Chemosensitive neurons in close apposition to large arteries could monitor rapid changes in perivascular CO2/Hþ due to fast changes in medullary arterial PCO2 and/or blood flow, provided they have the appropriate machinery for fast monitoring. Apart from a possible chemosensitive role in respiration, however, both the described cholinergic neurons in ventrolateral medulla [16] and the serotonergic cells in raphe [18] may have an excellent strategic position to play a modulating role in the adaptation of vessel caliber to the PCO2 and PO2 of the inflowing arterial blood. Note that also, hypercapnia-induced cerebral vasodilatation is based on a chemosensitive process governed by changes in local PCO2/Hþ.
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C. Searching the Anatomical Location of Central Chemoreceptors
Over the last decade, several new approaches were applied to localize the central chemoreceptors. This has contributed to a growing consensus to a widespread location of CO2 chemoreceptors in the brain stem. Currently, the following brain stem regions are assumed to contain CO2 chemoreceptors [32–39]: Dorsally, the Nucleus of the Solitary Tract (NTS) in the same areas where afferents from the carotid bodies project; the Locus Coeruleus (LC) in the pons; Ventrally (in rostrocaudal order), the RostroVentroLateral Medulla (RVLM) particularly the RetroTrapezoid Nucleus (RTN), Raphe Nuclei (RN) in the Rostral Ventral Medial Medulla (RVMM), the pre-Bo¨tzinger complex and a region within the Nucleus Ambiguous (NA) belonging to the group of respiratory neurons in the caudal ventrolateral lateral medulla. Some of these areas were explored with different techniques yielding corresponding results. Many other areas, however, may also contain chemoreceptors but are not as extensively explored (e.g., parabrachial and Ko¨lliker Fuse nuclei in pons and regions within the caudal mesencephalon—see below). In the following paragraphs we will discuss the most important tools that were (and still are) utilized to localize these many chemosensitive areas. Electrophysiological Studies In Vitro
In the last decade, the first evidence that several brain regions outside the ventral medullary surface contain CO2/Hþ chemoreceptors came from in vitro studies. In the neonatal rat spinal cord-brainstem preparation at depths between 50 and 700 mm from the ventral surface, respiratory neurons possessing extensive dendrites to within 50 mm from the surface were discovered, showing inherent CO2 sensitivity in the presence of synaptic blockade [13]. This preparation also contains CO2 sensitive neurons in the locus coeruleus [32,33]. In coronal brain slices, CO2/Hþ sensitive cells activated independently from synaptic events were described in the NTS [34,35], hypothalamus [36], locus coeruleus [37] and in the ventral medial medulla (raphe, 14). Intracellular recordings in ventral medullary raphe showed the existence of two chemosensitive cell types, one stimulated and one inhibited by CO2 [14]. This was confirmed by perforated patch recordings in neurons in cell cultures from isolated ventromedial medullary cells [38]. In addition, all acidosis-stimulated cells from these cultures appeared to be serotonergic and had a morphological appearance that clearly differed from that of the acidosis-inhibited cells that were all non-serotonergic [38,39]. Both types of acid-sensitive neurons showed a high sensitivity to pH changes in the physiological range [38].
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Teppema and Dahan Focal Injections of Acetazolamide
An unresolved issue is whether central chemoreceptors contain carbonic anhydrase, shown to be the case for the peripheral chemoreceptors in the carotid bodies [40–43]. Obviously, the presence of this enzyme in cells that should rapidly respond to changes in intracellular pCO2 would be useful. Unfortunately, however, this would not make it easier to localize and identify neurons that specifically function as CO2 chemoreceptors: like in other organs, carbonic anhydrase is a ubiquitous enzyme in the brain and has also been demonstrated in regions (e.g., hippocampal and cerebellar) that do not contain chemoreceptors (see below). Despite these limitations, local application of acetazolamide has proven to be a useful tool to localize chemosensitive regions in the brain stem, mostly in anesthetized animals. In several regions, namely NTS, RTN, midline raphe, rostral part of the ventral respiratory group, pre-Bo¨tzinger complex, and even fastigial nucleus in the cerebellum, topical application of acetazolamide is followed by increases in phrenic activity as large as 49–69% of the increase observed during overall hypercapnia following a change in end-tidal CO2 from 4 to 9% [44–49]. The injections cause acidotic responses in different focal chemoreceptor regions that, at the center of the injection sites, are equivalent to those following a change in end-tidal CO2 from 28 to 64–75 Torr [44–47]. In evaluating the effects of focal acetazolamide injections, the following points need attention: a. Local acidosis in putative chemoreceptor areas caused by acetazolamide injection does not always result in an increase in ventilation (e.g., 47). The reverse, a pronounced effect in chemosensitive regions (e.g., in midline raphe) known to contain relatively few carbonic anhydrase-containing chemosensitive neurons [50], has also been reported [46]. b. The mechanism by which acetazolamide injection causes focal acidosis is obscure and will remain so until more detailed information becomes available on the resulting effect on intracellular pH and on the identity, distribution and subcellular location of the (blocked) isoenzymes involved. One possible mechanism by which acetazolamide might cause extracellular acidosis could be by inhibiting a membrane-associated isoenzyme attached to neuronal processes which modulates the pH of extracellular fluid [51]. Inhibition of (intracellular) glial CA could also cause extracellular acidosis and this, together with a close association of glial cells with neurons, might explain why, as mentioned above, microinjection of acetazolamide in raphe increases respiratory output although this region possesses relatively few CA-containing chemosensitive neurons.
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c. The effects of rapidly permeating carbonic anhydrase inhibitors such as methazolamide were not systematically compared with those of an impermeable inhibitor such as benzolamide (note that acetazolamide is a slowly permeating inhibitor). The only study in which the effects of acetazolamide and methazolamide were compared showed a faster effect of the latter on phrenic activity suggesting involvement of an intracellular isoenzyme [48]. Another interesting feature of the latter study is that these inhibitors were injected into the pre-Botzinger complex, a region known to contain important respiratory neurons in rhythm generation [52]. Are these respiratory neurons chemosensitive? d. The time courses of changes in phrenic activity and local pH do not match: fast acidosis and gradually developing, long-lasting increases in ventilation [44,47]. This may be related to slow diffusion and permeability characteristics of acetazolamide, although a contribution of slow neuronal dynamics cannot be excluded. It is possible that the time course of the change in phrenic activity would match more closely to the (unknown) time course of the (acid?) change in intracellular pH. In this context it is interesting to draw a parallel with the peripheral chemoreceptors. In type I carotid body cells, acetazolamide causes alkalosis [53] and reduces the output and sensitivity of the carotid bodies [54–56]. This implies that on type I cells, carbonic anhydrase has an acidifying effect, which could be mediated by an extracellular membrane-bound isoenzyme that operates in concert with an HCO pump (bicarbonate out, chloride in; 53). This is an 3 /Cl unlikely scenario for the central chemoreceptors because acetazolamide applied locally [47] or via the cerebrospinal fluid [57] and methazolamide, administered systemically after prior inhibition of carbonic anhydrase in all peripheral tissues [30], all cause an increase in respiratory output and CO2 sensitivity. Full understanding of the role of carbonic anhydrase in central chemoreceptors will not be possible without detailed knowledge of the entire machinery that these cells possess to control intracellular pH: ion pumps (including the direction into which they operate), identity and subcellular distribution of carbonic anhydrase isoforms, possible interaction with glial cells, etc. e. Non-specific and/or other pharmacological actions of acetazolamide other than inhibition of CA alone cannot be excluded. For example, in guinea pig mesenteric arteries and human forearm vessels, acetazolamide seems to stimulate KCa (Maxi-K) channels, possibly by a specific action [58,59]. A similar effect may also be responsible for the reduction in O2 and CO2 sensitivity (and their interaction) of the peripheral chemoreflex loop by
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Microdialysis of CO2
Local application of CO2 in vivo with the aid of a CO2 microdiffusion pipette is a technique developed by Nattie and co-workers to examine the effect of local acidosis by a more physiologic stimulus than acetazolamide [61]. The design of the pipette tip is such that the tissue fluid injection can be avoided; the local acidosis is caused by diffusion of CO2 from the CO2-enriched artificial CSF flowing through the pipette. In contrast to the effects of local acetazolamide injections, the resulting acidotic stimuli rise and reverse quickly and have relatively short-lasting effects on ventilation. Thus, within one animal repeated ‘injections’ of CO2 can be made and dose-response relations studied. Similar to acetazolamide and dependent on the arousal state (anesthetized, asleep or awake), local application of CO2 causes large increases in ventilation when applied in caudal NTS, raphe and RTN, indicating the presence of CO 2 chemoreceptors in these regions. Focal dialysis in RTN with CSF equilibrated with 25% CO2, for example, caused a ventilatory response equivalent to that induced by a rise in arterial PCO2 by 1 kPa (about a 25% increase in ventilation; 61). In particular, application of this technique in awake and sleeping animals (rats) has revealed some other interesting properties of different chemosensitive regions. Microdialysis of CO2 in medullary raphe causes an increase in breathing (frequency) during sleep but not wakefulness [62], but the reverse was found for microdialysis in RTN, where an increase in tidal volume was found in the awake state only [63,64]. Acidosis in caudal NTS causes an increase in tidal volume and frequency while both awake and asleep [65]. Although it cannot be excluded that the same CO2 concentration in the pipette may cause different acidotic stimuli or stimulus spread in the awake state and during sleep, these results indicate that different chemoreceptor areas may have distinct roles depending on the arousal state. Carbonic Anhydrase Immunohistochemistry
In the brain, carbonic anhydrase is widely distributed. The most ubiquitous isoenzyme is CA II, which is found in the cytoplasm of oligodendroglia, microglia, choroid epithelium, in astrocytes and some neurons [66]. Because carbonic anhydrase-containing oligodendrocytes occur throughout the brain, it has been suggested that the enzyme plays an important role in
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controlling the excitability of neurons by rapidly converting metabolic CO2 to Hþ [67]. Membrane-bound CA II has also been demonstrated and constitutes about 50% of the total CA activity in rodent brains [68]. Another isoenzyme, CA IV, a membrane-associated isoform, has been localized both in capillary endothelium, glia and myelin and to a lesser degree in neurons [69–72]. Generally, with the exception of the rostromedial and rostroventrolateral medulla, a detailed topographic description of CA isoenzymes in chemosensitive areas is not available. In rostroventrolateral medullary areas associated with central chemoreceptors and in chemosensitive neurons cultured in vitro, cytosolic and membrane-bound isoenzymes have been demonstrated [73,74]. Wang and co-workers [38] showed that within the medullary raphe many serotonergic neurons do not show immunoreactivity to CA II and/or CA IV. Furthermore, they showed that many other neurons, e.g., hippocampal or cerebellar, do stain for CA, although these regions contain only small percentages of chemosensitive neurons. They also showed examples of serotonergic raphe neurons cultured in vitro (not stained for CA) which were highly sensitive to CO2 but insensitive to acetazolamide. The above data indicate that CA cannot be considered a unique marker for chemoreceptors. Why do some chemoreceptors contain some isoform of the enzyme while others do not? We suggest that this may be related to their location relative to blood vessels. If the arterial PCO2 rises upon exposure to CO2, chemoreceptors that are closely associated with larger, superficial arteries will be more rapidly subjected to a rise in perivascular PCO2 than those supplied with blood from smaller vessels further downstream. Local presence of CA could enable these cells to translate these alterations rapidly into changes in intracellular pH. In this context we refer to an older study in carotid body denervated cats in which we found that changing the extracellular pH in the medulla by way of exposing the animal to squarewave changes in end-tidal CO2 led to a change in phrenic activity that could be well described with a model (with the extracellular pH as input) containing both phasic and tonic components [29]. In this study, the extracellular fluid (ECF) pH changed with a time constant of about 40 s. Identification of a fast and slow component, possibly representing fast and slow chemoreceptors, would be easier if it were possible to induce squarewave changes in medullary ECF pH. Due to (perfusion-limited) slow washin or wash-out of CO2, this is very difficult. It would be interesting, however, to investigate if the distribution of carbonic anhydrase within chemosensitive neurons may be related to their proximity to blood vessels. For example, double staining for Fos and carbonic anhydrase in animals exposed to CO2 breathing could tell us if a preferential staining for carbonic anhydrase may occur in activated (Fos-positive) neurons with cell bodies or processes closely associated with vessels.
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Teppema and Dahan c-fos Immunohistochemistry–A Tool to Localize Central CO2 Chemoreceptors?
The nuclear proto-oncogene c-fos is an immediate early gene that may link short-term cytoplasmic and membrane responses with adaptive alterations in gene activity resulting in both short- and long-term changes in neuronal function [75]. When a cell is exposed to a stimulus, a second messenger causes the induction of immediate-early genes, which then is followed by synthesis of protein (complexes) that function in signal transduction [76,77]. Thus, proto-oncogenes can encode ligands, receptors, GTP-binding proteins, tyrosine-kinases, nuclear receptors and DNA-binding proteins. Expression of c-fos is considered as a marker of activation of individual neurons after synaptic activation and influx of Ca2þ through voltagesensitive Ca2þ channels [76–79]. The protein product of c-fos, the nuclear protein Fos, is rapidly and transiently induced and remains in the nucleus for several hours, where it can be identified with the aid of specific antibodies. Initially, in respiratory physiology, Fos immunohistochemistry was presented as a tool to identify central chemoreceptors [80], and many investigators have used the technique to map neuronal pathways in the brain that are activated during hypercapnia (and also hypoxia). Here we give a brief overview of the most important sites that appear to contain increased Fos levels after exposure of intact animals to inhalation of CO2 (nomenclature after [81]; data from [80,82–91]): a. in dorsal medulla: the commissural, dorsomedial and medial subnuclei of the Nucleus Tractus Solitarius (NTS) at caudal level and at the obex, with little staining in the ventrolateral subnuclei; b. in the Caudal Ventrolateral Medulla (CVLM): A1 noradrenergic cells, but also non-catecholaminergic neurons within and ventral to the Lateral Reticular Nucleus (LRN) and in the vicinity of the nucleus (retro)ambiguous; c. in the Rostral Ventral Medial Medulla (RVMM): neurons in raphe pallidus (RP) and magnus (RM) and gigantocellular nucleus pars a (GiA); d. in Rostral VentroLateral Medulla (RVLM): many neurons within the area overlapping the A1 (noradrenergic) and C1 (adrenergic) regions, within the C1 region, also called the subretrofacial area, retrofacial lateral ParaGigantoCellular nucleus (PGCl) and rostral ventrolateral nucleus [84]. The ventral border of this area is analogous to the intermediate chemosensitive area in the cat [7]. At the rostral border of the RVLM, just caudally from (and in the cat ventrally to) the facial nucleus many c-fos expressing cells are present in a region called the RetroTrapezoid Nucleus (RTN) (Figure 2.2, taken from the cat medulla);
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Figure 2.2 Activation of RTN (retrotrapezoid nucleus) cells in the rostroventrolateral medulla of an anesthetized cat that was exposed to 10% CO2 for 1 hr. Black spots are nuclei that stained for the protein Fos which is the product of the immediate early gene c-fos (Data from Ref. 82).
e. at pontine level: within the Locus Coeruleus (LC), Kolliker-Fuse nucleus and A5 noradrenergic cell group, and more rostrally the external subnucleus of the parabrachial nucleus (vlPAG); f. in caudal mesencephalon: the ventrolateral periaqueductal gray and dorsal raphe structures; g. in more rostral brain areas: the medial supramammillary nucleus, the central nucleus of the amygdala, the paraventricular and supraoptic nucleus in the hypothalamus, the medial preoptic nucleus, the bed nucleus of the striae terminalis, and the lateral septum nucleus; h. in one study, expression of c-fos was reported to occur in rostral regions of the fastigial nucleus in the cerebellum [92]. The above data come from studies showing considerable differences in outcomes, which is not surprising for several reasons. First, the time pattern of Fos expression strongly depends on the strength, duration and nature of the stimulus. Second, the precise experimental paradigm may be of utmost importance because the Fos protein acts as a negative regulator of its own expression: after an initial stimulation (for example, a short period of hypercapnia preceding the test period), there may be a refractory period for a re-induction of c-fos expression that may last for several hours [75]. Thus
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studies with different experimental paradigms may be expected to yield different results. Third, variations in perfusion, fixation, and preparation techniques as well as in antibody specificity or dilution may significantly influence the staining results. In addition, some studies were performed in awake animals, while in others an anesthetic preparation was used. Various anesthetics influence the expression of c-fos differently [93–95]. Anesthesia may also alter hypercapnia-induced expression of Fos in some regions. For example, hypercapnia induces c-fos expression in LC and medial preoptic nucleus in awake rats but not in those anesthetized with a-chloralose urethane [96]. An additional source of confusion may be born from the fact that many Fos-expressing neurons are located in the reticular formation of the brainstem where clearly defined functional regions with wellcircumscribed structures and boundaries are absent (some authors use coarse rather than precise anatomical descriptions). Apart from the above factors, a few other factors should be taken into account. First, hypercapnia is not a specific stimulus to the ventilatory control system because it also elicits cardiovascular, sympathetic and neuroendocrine responses. Second, expression of c-fos is not limited to specific cells but may play a role in stimulus-response coupling that is common to most cell types. Multiple second-messenger pathways appear to converge on c-fos, which is compatible with the finding that many cell types express the gene [75]. In this context, it is of interest to note that there is considerable overlap of brain(stem) regions showing increased expression of c-fos during both hypercapnia and hypoxia [84,88–90,97]. Brain stem regions of the rat showing increased expression during hypercapnia but not hypoxia are RTN, juxtafacial PGCl, GiA, LC and vlPAG [84]. Finally, expression of c-fos may be dissociated from neuronal firing; in some cases increased expression may mirror an increased level of second messengers rather than neuronal activity [75,98]. On the other hand, despite the ability of many cells to express c-fos, the absence of increased Fos levels does not prove absence of activation, so that negative outcomes cannot be excluded. The above limitations and uncertainties may give the impression that using Fos immunohistochemistry raises more questions than it answers. It does not mean, however, that it could not be a useful tool in the identification of central chemoreceptors. First, there is fair agreement between the location of neurons in the brain stem with increased Fos levels after a hypercapnic challenge and those chemosensitive cells that are localized with microinjection of CO2 and acetazolamide and with electrophysiological means. This is particularly true for neurons in the NTS, CVLM, raphe structures in the RVMM, subretrofacial area and RTN in the RVLM and locus coeruleus in the pons. Figure 2.3 shows similar locations of chemosensitive neurons in ventral raphe of the rat medulla identified with electrophysiological means (left panel), and those of
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Acidosis-stimulated Acidosis-inhibited
LPG
B
Pyr
0.25 mm
Figure 2.3 Chemosensitive cells in rat medullary raphe localized with electrophysiological means in vitro (left panel, data from Ref. 15) and with Fos immunohistochemistry after exposing the whole animal to 15% CO2 for 1 hr (right panel, data from Ref. 84). B: basilar artery; Pyr: pyramidal tract; LPG: lateral paragigantocellular nucleus.
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Figure 2.4 Expression of Fos in the external lateral subnucleus of the parabrachial nucleus of a rat exposed to 10% CO2 for 1 hr. Whether the stained nuclei are from chemosensitive cells is unknown (Data from Ref. 84).
Fos-positive cells after exposure to CO2 in vivo (right panel). Other regions containing immunostained and possibly chemosensitive cells, cells such as the parabrachial nucleus in the pons (Figure 2.4) and more rostral regions in the hypothalamus are relatively unexplored with these alternative techniques, with the exception of electrophysiological studies showing CO2 responsive neurons in the caudal hypothalamus [36]. Note also that Fos immunohistochemistry is a method to localize somata (nuclei) of activated neurons and does not provide any information about the location of the possibly distant chemosensitive sites. Fos immunohistochemistry is a technique with single-cell resolution, a clear advantage. Thus it has an enormous potential to characterize individual activated neurons neurochemically, provided Fos staining is combined with the use of specific antibodies directed against neuromediators, transmitters, second messengers and enzymes. In this way, for example, we know that the majority of neurons in the RVLM that are activated during hypercapnia do not contain noradrenaline or dopamine [84]. By using antero- and retrograde tracers, functional properties of neuronal circuitries can be studied in different physiological circumstances. Fos immunohistochemistry can also be used in combination with pharmacological tools, for example to reveal the second messenger associated with hypercapniainduced Fos expression or to investigate the causal relationship between the increase in ventilation and the increased expression (e.g., by using anti-sense Fos mRNA).
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LHB LPB VMH
SON OT
LC
DM
AP NTS
Pons
7
PreBotc cVRG PGCL VRG 10
rVMS RTN
BotC
P
cVMS
Figure 2.5 Fos expression in the rat brain in response to stimulation of the rostral chemosensitive area at the brain stem surface of the rat with acidic CSF (pH 7.2). CVRG: caudal ventral respiratory group; DM: dorsomedial hypothalamus; 10: inferior olive; LC: locus coeruleus; LHB: lateral habenular nucleus; LPB: lateral parabrachial nucleus; NTS: nucleus tractus solitarius; PGCL: lateral paragigantocellular nucleus; preBotc: pre-Bo¨tziger complex; RTN: retrotrapezoid nucleus; 7: facial nerve; VMH: ventromedial hypothalamus; SON: supraoptic nucleus; OT: optic tract; rVMS: rostral ventral medullary surface; cVMS: caudal ventral medullary surface; P: pyramidal tract. AP: area postrema (Data from Ref. 99).
In summary, more insight is needed into the mechanism(s) by which so many neurons in the brain stem (and more rostral brain regions) show increased expression of c-fos during hypercapnia: are all these neurons directly activated by CO2 or are synaptic events involved? In this context, two recent studies are of particular interest. Douglas et al. [99] showed increased expression of c-fos in many neurons in brainstem, hypothalamus and more rostral regions after application to the caudal and rostral ventral chemosensitive areas of pledgets soaked in artificial CSF with an acidic pH of 7.2. This shows that exposing the ventral medullary surface areas to acid initiates c-fos induction in many distant areas (Figure 2.5), several of which are also activated during exposure to CO2 of the whole animal. Some of these neurons may be activated because they possess (long) processes projecting to the ventral surface. Other neurons, however, may have been activated transsynaptically, as the caudal and rostral ventrolateral medulla have many connections with other brain(stem) areas [84]. Okada et al. [16] studied the expression of c-fos in the ventral medulla after in situ transarterial perfusion of the animal with mock CSF equilibrated with
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8% CO2 and containing low calcium, high magnesium or TTX to block synaptic transmission. The ventral medulla of animals perfused with normal CSF (also equilibrated with 8% CO2) without the synaptic blockers contained many stained large- and small-size nuclei in cells located predominantly in perivascular superficial tissue. In animals that were perfused with CSF containing the synaptic blockers, Fos-containing cells (mostly with small nuclei) were restricted to the marginal glial layer close to the medullary surface and closely associated with blood vessels, suggesting that they may represent true CO2 chemoreceptors. Indeed, in a perforated patch-clamp configuration, the same superficial region appeared to contain cells that were activated by hypercapnia. These well-designed studies show that Fos immunohistochemistry can be a very valuable tool in the identification of central CO2 chemoreceptors.
III.
Mechanism of Central Chemoreception
A. Extracellular pH, Intracellular pH, CO2, and Central Chemoreception
Several in vivo data indicate that the intracellular (pHi) rather than the extracellular pH (pHe) plays a crucial role in central chemoreception. In the anesthetized cat, isocapnic medullary pHe changes have a much smaller effect on minute ventilation or phrenic output than equal changes induced by hypercapnia [9–11]. This can be explained by a major role of pHi in chemoreception or, alternatively, by a direct effect of CO2. After systemic administration of acetazolamide in carotid body denervated cats, the time course of the resulting changes in medullary pHe and ventilation do not match [29], indicating that another factor, possibly pHi, acts as stimulus. The same applies to local application of acetazolamide into chemosensitive areas of rat and cat medulla [44,47]. Intracellular acidosis in the ventral medulla, however, does not always increase ventilation. After peripheral chemodenervation, a hypoxia-induced acidification of the ventral medullary surface fails to increase ventilation in anesthetized rabbit and cat [100,101]. The same was found in awake goats undergoing CO-hypoxia [102]. Assuming intracellular acidosis as the source of this extracellular acidosis, it was suggested that the absence of an increase in ventilation could be explained by an increase in the intra- to extracellular Hþ gradient [100–102]. Hypercapnia or a metabolic acidosis originating in the extracellular environment would decrease this gradient and hence stimulate the chemoreceptors. Organotypic medullary cultures in which many features of central chemoreceptors are preserved are successfully used to study chemoreceptor pHi and pHe responses to manipulations such as hypercapnia (causing both
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intra- and extracellular acidosis), replacement of bicarbonate by HEPES in the bath solution (causing a fall in pHi but no change in pHe), ammonia wash-in (extracellular acidosis, intracellular alkalosis) and ammonia washout (extracellular alkalosis, intracellular acidosis; [103]). In all cases, neuronal responses correlate with changes in pHi rather than pHe, with a fall in pHi consistently coupled to an increase in activity [103,104]. Another argument for pHi as the proximate signal for central chemoreception has been the observed steeper pHi–pHe relationship during hypercapnia in neonatal chemosensitive neurons compared with nonchemosensitive cells [105,106]. Most nonchemosensitive neurons in the inferior olive and hypoglossal nucleus show pHi recovery upon exposure to a hypercapnic acidotic solution; chemosensitive neurons from the ventrolateral medulla and NTS, however, fail to show this recovery, except when the intracellular acidosis is caused by an isohydric (i.e., constant pHe) hypercapnia [105,106]. Carotid body type I cells [53] and taste-bud receptor cells [107] show a similar behavior, suggesting that it might be a common feature of CO2-sensitive cells. It has been suggested that this failure of chemosensitive neurons to recover may be caused by inhibition of a Naþ/Hþ exchanger (NHE, acting as an acid extruser and located in the chemoreceptor membrane) by the low extracellular pH. Naþ/Hþ exchangers in the membrane of chemoreceptors would be much more sensitive to changes in pHe than those located in the membrane of nonchemosensitive cells, explaining a much steeper pHi–pHe relationship in the former. Similarly, a high sensitivity of the anion (Cl/HCO 3 ) exchanger of ventral medullary chemoreceptor cells might explain the failure of these cells to show recovery upon an alkaline challenge [105,106,108]. The cause of this difference in pHe sensitivity of chemoreceptor vs. nonchemoreceptor NHE is unknown, but it has been related to the presence of a different isoform of the transporter in the chemoreceptor membrane. From the six NHE subtypes known to occur in the brain [103], the isoform NHE3 seems to be preferably located in neurons of the superficial ventrolateral region of the medulla [103,109]. Also, selective NHE3 inhibitors are reported to mimic CO2 responses of chemosensitive neurons in vitro, to reduce pHi in VLM neurons and to inhibit pHi regulation upon exposure to ammonia [103]. In the anesthetized cat and rabbit, systemically administered selective NHE3 inhibitors reduce the apneic threshold (own unpublished observations; [110]). The idea that central chemoreceptors may differ from other neurons in their pHi regulation machinery for which a specific membrane transporter might be responsible could be useful in future attempts to isolate unique features of these chemoreceptors. It should be noted, however, that the above data showing pHi regulation in nonchemosensitive cells come from neonatal and therefore immature animals. Nottingham et al. [111]
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compared pHi responses in RTN and hypoglossal neurons from neonatal and mature animals. While neonatal hypoglossal but not RTN neurons showed pHi recovery upon a hypercapnic exposure, in cells from mature animals this difference disappeared: both mature cell types showed no pHi recovery, unless amiloride (to inhibit NHE) was added to the bath. Thus the question remains, whether in mature chemosensitive cells a lack of pHi recovery from intra- and extracellular acidosis indeed represents a feature of central chemosensitivity rather than a general mechanism to protect neurons against an NHE-induced influx of Naþ ions which eventually may lead to cell death [111]. Because hypercapnic exposure with a constant pHe results in intracellular acidosis and increased neuronal activity, it is reasonable to ascribe an important role to the intracellular pH in central chemoreception. A steep pHi –pHe relationship in hypercapnia will imply an appreciable sensitivity to changes in pHe under this condition. Because the relation between pHi and pHe (and vice versa) during a given change in acid-base status depends on the origin of the primary disturbance, it is difficult to describe the relation between ventilation (or chemoreceptor activity) and pHe in terms that are generally valid. A further reason for the complex ventilation-pHe relation is that the extracellular pH may influence chemoreceptor activity in a number of ways independently from its effect on pHi. Various types of membrane ion channels, for example, may change their conductance upon pHe changes and initiate an intracellular chemosensitive stimulus-transduction cascade (see below). Changes in pHe may also modulate ongoing synaptic transmission at the chemoreceptor membrane. A notable and possibly important example is the known pH sensitivity of several members of the purinergic receptor family [112]. In the anesthetized rat, microinjection of the selective P2X antagonist suramin in the retrofacial area increased the apneic threshold and reduced the respiratory response to CO2 [112]. An even more interesting observation was that suramin and PPADS (pyridoxal-phosphate-6-azophenyl-20 ,40 disulfonic acid, another P2X receptor antagonist) reduced the ongoing activity of inspiratory and pre-inspiratory neurons in the pre-Bo¨tzinger complex and blocked their activation by CO2 and ATP [113]. If indeed the excitation of these respiratory neurons by ATP and the P2X agonist ab-methyleneATP would involve a pH-sensitive purinergic receptor, then this would be another clear example of respiratory neurons showing direct pH sensitivity [also 12,13]. Cholinergic neurotransmission is a second example of a pHe-modulated transmission. From numerous studies it is clear that, at least in the ventral medulla, the central chemosensitive mechanism involves a muscarinic cholinergic mechanism [114]. In the in vitro brainstem-spinal cord preparation, blockage of M1 and M3 muscarinic receptors resulted in a depression (cessation) of respiratory output, an effect that was easily
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reversed by facilitation of acetylcholine release [115]. In the cat, topical application of atropine to the ventral medullary surface blunted the ventilatory response to CO2 [116]; this effect could be reproduced by using M1 and M3 receptor blockers [117,118]. In the unanesthetized rat and newborn piglet, microdialysis of muscimol in the RTN inhibited the response to systemic hypercapnia by 24% and 40%, respectively [119,120]. In slices from ventral medulla of rat, Fukuda and Loeschcke [121] showed that an intact cholinergic transmission was a prerequisite for activation by acidic solutions. A similar observation was made in spontaneously breathing dogs undergoing ventriculocisternal perfusion: an increase in ventilation in these animals by CSF acidification was abolished in the presence of atropine [122]. What is the link between chemosensitivity, cholinergic transmission and pHe? The acid sensitivity of cholinergic neurotransmission could reside in a pH effect on imidazole groups of the hydrolytic enzyme acetylcholinesterase [114]. From imidazole groups, on the other hand, we know that they are involved in central chemoreception because when applied to the ventral surface of cats, imidazole-binding agents inhibit the ventilatory response to CO2 [123; for a further explanation of the a-stat hypothesis of central chemoreception see 124]. The varying pHi –pHe relation on one hand, and the additional independent effects of extracellular pH on the other, make the relation between chemoreceptor activity and pHi extremely complex. For example, during brain stem hypoxia the chemoreceptor NHE may be activated [125,126], tending to recover pHi from intracellular acidosis. The resulting (actually measured) fall in pHe may now be unable to inhibit the (activated) membrane transporter. If we assume a sustained intracellular acidosis in the case of steady-state hypoxia (to which increased uptake of lactate via a monocarboxylate transporter may also contribute [127]), then the intriguing question remains about what happens to central chemoreceptor output in this situation: is the failure of minute ventilation to increase (see above) due to independent effects of pHi and pHe on the chemoreceptor neuron, or rather to effects unrelated to the mechanism of chemoreception such as inhibitory modulators acting elsewhere on the respiratory network? It may prove to be elusive to isolate a unique stimulus for the central chemoreceptors, because there are many ways to depolarize a neuron (apart from the question whether depolarization is a conditio sine qua non for activation; see below). A given change in pHe or pHi may have entirely different effects on individual chemoreceptor cells depending on their membrane properties and location (see also below). Should neurons that depolarize upon a fall in pHe, despite the fact that they may not possess the specific machinery of chemosensitive cells in VLM and NTS to regulate pHi, also be called central chemoreceptors? (cf. hypoglossal motoneurons that have TASK-1 channels in their membranes and
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respond with depolarization upon acidosis; see below). Finally, although direct effects of CO2 cannot be excluded, it also cannot be considered as the adequate stimulus because there are many data showing that isocapnic changes in pHe and pHi alter the activity of chemosensitive cells. B. Intracellular pH, Gap Junctions, and Central Chemoreception
One possible explanation for the surprisingly large effects of topical application of CO2 and acetazolamide in chemosensitive brainstem areas on ventilation (see above) is that focal stimulation is spread out over an entire network consisting of neurons that can read an electrical (or metabolic) signal generated by the membrane of a neighboring cell via gap junctions. Gap junctions form a low-resistance communication pathway between adjacent cells and are made from two hemi-channels (connexons), each of which consists of a hexamer of connexins (Cx) [128]. Immunohistochemical studies have shown that putative chemosensitive areas in the NTS, retrofacial area and RTN, but also in the pre-Bo¨tzinger complex, contain gap junction proteins: Cx26 in neurons and astrocytes and Cx32 in neurons [129,130]. This suggests the presence of gap junctions in these areas via which central chemoreceptors could be electrotonically coupled. Indeed, a large percentage of CO2-sensitive neurons in NTS appears to be anatomically and chemically coupled, in contrast to most cells that do not respond to CO2 [131,132]. Many CO2-sensitive neurons in LC also show cell-cell coupling [33,131,132]. Gap junctions appear to close in response to a rise in intracellular [Ca2þ] or [Hþ] [133–135]. Because the central chemoreceptors most likely respond to changes in intracellular pH, closing of gap junctions in response to a rise in intracellular [Hþ] could play a role in the chemosensitive mechanism. Interestingly, Cx32, one of the gap junction proteins in the presumptive chemosensitive areas in the ventral medulla, contains a cytoplasmic domain that is critical to CO2/pH gating sensitivity [136,137]. However, spontaneous electrotonic postsynaptic potentials of LC neurons do not seem to be affected by hypercapnic acidosis, and synchronization in pairs of LC neurons does not disappear with acidosis in the physiological pH range [131,138–140]. This may be related to the fact that the coupling coefficient between coupled neurons also depends on non-junctional conductance [128]. Thus, a decrease in nonjunctional (membrane) conductance in chemosensitive cells, for example during an acidosis-induced depolarization, may tend to enhance junctional cell coupling and offset the channel-closing effect of the lower intracellular pH. As mentioned above, gap junctions are sensitive to intracellular pH. After uncoupling, however, chemosensitive neurons in the solitary complex
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and locus coeruleus retain their CO 2 sensitivity, indicating that cell-cell coupling is not a requirement for CO2 sensitivity per se [33,131]. Coupling between cells resulting in synchronization may have considerable effect on the gain of a system [128]. This may be one of the factors that determine the CO2 sensitivity of the entire network of central chemoreceptors or the form in which CO2 sensitivity is expressed, e.g., DC (low frequency) changes in membrane potential vs. changes in spike frequency [33]. Another function of gap junctions in chemoreceptor cells may be related to the exchange of second messengers such as Ca2þ, cAMP and IP3, allowing coregulation of these messengers in pre- and postsynaptic compartments and the modulation of presynaptic release by postsynaptic second messengers [128,141]. Future studies should address several important questions for more insight into the role of electrotonic and/or biochemical coupling in central CO2 chemoreception. First, it is of interest to document the particular (homocellular or heterocelllular) types of potential gap junctions within the ventrolateral medullary chemosensitive areas and their electrophysiological behavior. Second, what is the effect of changes in intracellular pH on cell coupling between chemoreceptor cells? Is the apparent lack of uncoupling by CO2 (low pH) unique to coupled cells that are excited by CO2? Third, what would be the influence of variations in the level of coupling between chemoreceptors on the CO2 sensitivity of individual neurons and on the gain of the entire network? Could various neurotransmitters (e.g., dopamine and serotonin, transmitters known to influence cell-cell coupling [128]), alter the coupling coefficient between chemoreceptor cells via an effect on membrane resistance and, by this, the sensitivity of the entire network? Finally, could the uncoupling effect of a volatile anesthetic such as halothane [128] be responsible for its depressant effect on central chemosensitivity? C. Glia, Extracellular pH and Central Chemoreception
It is well established that glial cells play a key role in the regulation of the ionic ([Hþ] and [Kþ] in particular) extracellular microenvironment of neurons, which means that the glial cells are a crucial element in determining the neurons’ stimulus level and excitability [142]. What does this mean for the central chemoreceptors? Glial cells could influence the central chemoreceptors in several ways; first, by regulating the extracellular pH [143] which at many locations occurs with the aid of carbonic anhydrase, providing a machinery for fast pHe regulation [66–68]. Local injection of the glial toxin flurocitrate into the RTN results in local extracellular acidosis and an increase in phrenic activity and spontaneous ventilation in anesthetized and conscious rats, respectively [144,145]. CO2 sensitivity is retained in these circumstances [145], indicating
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that the integrity of glial cells is not crucial for an intact chemosensitive mechanism per se. The ventilatory stimulation observed in the above studies may indeed be due to the extracellular acidosis, but a possible influence of a disturbed extracellular potassium concentration cannot be excluded. By regulating the extracellular [Kþ] [142,146], glial cells will directly influence the membrane potential and excitability of neurons. Synthesis, release and uptake of neurotransmitters is another way by which glial cells could influence the activity of central chemoreceptors or even the chemosensitive mechanism. For example, glial cells release lactate that is formed from glucose; this process is stimulated by glutamate that is released from neurons. The lactate then serves as a nutrient for the neuron [127]. Finally, the functional role of possible neuron-to-glial coupling or glial-glial coupling in the central chemosensitive network remains to be determined (see above). D. Intracellular pH, Extracellular pH, Membrane Channels, and Central Chemoreception
Even after synaptic blockade, chemosensitive neurons in NTS, raphe and RVLM respond to CO2/Hþ, suggesting that ion channels may be the primary responsive elements initiating an increase in spike frequency [13,14,147,148]. Dean et al. [148] found that CO2/Hþ depolarized NTS neurons via a reduction in potassium conductance of the membrane, without specifying particular candidates. The large family of potassium channels has many members that possess pH sensitivity. Notable examples are TASK (TWIK-related acid sensitive) channels. These two-pore-domain background potassium channels show fast activation and inactivation kinetics, and are extremely sensitive to changes in external pH within a narrow range (midpoint inhibition of TASK-2 ¼ pH 8.3, TASK-1 ¼ pH 7.3 and TASK-3 ¼ pH 6.3; [149,150]). TASK currents have been recorded in chemosensitive locus coeruleus and serotonergic raphe neurons, but also in hypoglossal motoneurons [151–153]. Thus, because inhibition of these currents in (chemosensitive) cells by extracellular acidosis in the physiological pH range results in their depolarization and increase in excitability, it is attractive to consider TASK channels (and particularly TASK-1) as potential primary targets for CO2/Hþ to initiate the chemosensory stimulus-transduction cascade. In type-I carotid body cells from neonatal rats, inhibition of TASK-1 channels by hypoxia and acidosis has also been shown to result in membrane depolarization [154]. It is of interest to note that in hypoglossal pH-sensitive motoneurons and cerebellar granule cells, TASK channels are modulated (inhibited) by various G-protein-coupled neurotransmitters that are also able to stimulate central chemoreceptors or respiratory neurons, e.g., acetylcholine, TRH, glutamate and serotonin [151,153,155].
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Transmitter Ligand Receptor 5-HT 5-HT2 5P NK1 TRH TRH-R1 α1 NE glutamate mGluR (group 1)
− Gαq-coupled receptors
TASK-1
+
K
−
+
Anesthetic
Figure 2.6 Modulation of Task-1 channel conductivity by pH, anesthetics and transmitters (Data from Ref. 153).
Also, opening of TASK channels is one of the known concentrationdependent effects of volatile anesthetics such as halothane and isoflurane [153,154,156–158]. This could provide a molecular basis not only for the anesthetic-induced decrease in excitability of motoneurons [153,158], but also for the known depressant effects of these agents on the central (but also peripheral) chemoreflex loop [153,154,159–162]. The above modulating influences on TASK-channel conductivity are summarized in Figure 2.6. Inhibition by extracellular acidosis of TASK channels cannot provide a complete scenario for central chemoreception simply because central chemoreceptors can be activated independently from changes in pHe. It also appears possible to increase spike frequency without membrane depolarization in locus coeruleus and medullary raphe neurons by applying an isohydric (i.e., constant pHe) hypercapnic stimulus [72,163]. Also, recent patch-clamp studies have shown that (hypercapnic) acidosis inhibits Ca2þactivated Kþ (KCa) channels in chemosensitive medullary raphe neurons and in fetal medullary neurons either by a direct effect on these channels or by inhibition of voltage-sensitive Ca2þ channels [14,164]. Note that KCa channels are inhibited by intracellular acidosis [165,166]. In locus coeruleus neurons from rat, isohydric and hypercapnic acidosis have been shown to stimulate L-type Ca2þ and multiple Kþ channels that are expressed in these neurons, e.g., KCa, TASK and inward rectifying Kþ channels (Kir) that is, similar to KCa, also show sensitivity to the intracellular pH [37,167]. The Kir channels deserve special mention because some members or heteromeric combinations are extremely sensitive to the intracellular pH, so that these channels could be a potential tool in the localization of central chemoreceptors. Jiang et al. [168] have presented a very elegant approach. Because it was already known that Kirs are involved in the control of membrane excitability and in cellular responses to hypercapnia and acidosis, these investigators used mammalian cell lines to express various homomeric Kirs and identified several subtypes that are sensitive to changes in
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intracellular (Kir1.1 and Kir4.1) and extracellular (Kir2.3) pH. Subsequently, they identified the critical pH-sensitive residues and motifs of these channels, and, after searching the GenBank, found an amino acid sequence in Kir5.1 identical to the critical motif of Kir2.3. Kir5.1 owes its biological significance to the fact that it forms heterodimeres with the Kir4 subfamily [168]. Because the heterodimer Kir4.1–5.1 showed a higher pH sensitivity than other members of the family and a pKa of 7.45, they suggested that it may well function at physiologic pH and PCO2 levels. When coexpressed in Xenopus oocytes, the current through these Kir4.1–5.1 complexes was reduced at high (8%) CO2 levels (leading to depolarization) but enhanced at low CO2 (3%) (leading to hyperpolarization), compared with the baseline current at 5% CO2. This led the authors to suggest that this Kir4.1–5.1 complex could enable cells to increase or decrease their membrane excitability in response to a rise or fall in PCO2, respectively. As a great surprise, using in situ hybridization, the authors then showed coexpression of Kir4.1–5.1 in cells belonging to regions proposed to contain central chemoreceptors: locus coeruleus, nucleus tractus solitarius, and within the caudal and rostral ventrolateral medulla (e.g., subretrofacial area). This well-designed strategy shows that in order to identify the central chemoreceptors, beginning with molecular mechanisms of CO2/Hþ chemoreception may be a very promising approach. E.
Conclusion
To date, there is no general model for the entire stimulus-transduction cascade of CO2 chemoreception. A general picture emerging from the available data is that individual chemoreceptors may display considerable differences in their pHi regulation machineries, receptors and ion-specific membrane channels that are sensitive to changes in either pHi, pHe or both. The fact that central chemoreceptors are so widely distributed over regions with different afferent connections containing specific transmitters and modulators adds to this complexity. Receptors that influence the conductance of ion channels and/or ion channels themselves (e.g., TASK) are molecular targets for these modulators; this may explain the effects on chemosensitivity of agents such as acetylcholine, ATP and other transmitters (e.g., glutamate) and their antagonists. Variations in tonic or phasic influences of these neuromodulators may thus lead to variations in the stimulus-transduction cascade in chemosensitive cells once they are stimulated by CO2/Hþ. All these factors make it an extremely difficult challenge to isolate a neurochemical or electrophysiological property that is unique for central CO2 chemoreceptors. There is growing consensus of a prominent role for the intracellular pH and potassium (but also other) channels during hypercapnia, but also of an
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important influence of the extracellular pH. Involvement of (acid-sensitive) potassium channels seems to fit in a general framework of CO2 chemoreception in which closure of these channels leads to depolarization and activation. Particularly, a significant role of TASK channels may emerge because they are sensitive not only for changes in extracellular pH but also for neuromodulators and anesthetics that are known to alter chemosensitivity.
IV.
Central Chemoreceptors and Breathing
The central chemoreceptors are an essential part of the control system regulating pulmonary ventilation. By providing a feedback signal representing local blood flow, metabolism and acid-base status of the body to the respiratory controller, they are a crucial factor in pH homeostasis of the body. Central chemoreceptors are the major structures responsible for the ventilatory response to CO2. They also provide a tonic drive to the respiratory motoneurons; especially during sleep, an intact central chemosensitive network seems crucial for the maintenance of an adequate level of ventilation that is important particularly during NREM sleep. The widespread distribution of central CO2 chemoreceptors throughout the brainstem suggests that they may participate in various specific functions associated with the control of breathing but also in functions associated with the control of the arousal state. All these aspect will be briefly discussed in the following section. A. Central Chemoreceptors and Ventilatory Response to CO2
The exact quantitative contribution of the central chemoreceptors to the ventilatory response to CO2 is somewhat controversial, although there is general consensus about a central contribution of at least 50–60%. The issue has been addressed with various approaches, discussed briefly below. Ventriculocisternal Perfusion
In their classic ventriculocisternal perfusion experiments, Fencl et al. [3] attributed the ventilatory adaptations to metabolic and respiratory acidbase disturbances entirely to central chemoreceptors functionally located at two-thirds of the distance between cerebrospinal fluid and arterial blood. In their calculations, the authors treated the extracellular pH as the adequate chemoreceptor stimulus but later findings by other groups were in disagreement with this assumption (e.g., [9–11]). Furthermore, by neglecting the influence of the carotid bodies in hyperoxia, the role of the central chemoreceptors in this preparation was overestimated.
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From dynamic CO2 studies in man under hypoxic, normoxic and hyperoxic conditions, it was initially concluded that hyperoxia virtually eliminates the contribution of the peripheral chemoreceptors to the CO2 response, and thus would provide a means to study central CO2 sensitivity independently of the peripheral chemoreceptors [169,170]. However, now there is ample experimental evidence to show that this assumption overestimated the role of the central chemoreceptors, because even during hyperoxia the carotid bodies may possess considerable CO2 sensitivity. In the anesthetized cat, for example, hyperoxia does not eliminate the CO2 sensitivity of afferent carotid sinus nerve afferents [171–173]. This may be a species-dependent phenomenon, however, because in the anesthetized rabbit, the carotid bodies are virtually silent at PO2 values 430 kPa [174]. Similar to the cat, humans show a considerable peripheral CO2 sensitivity during hyperoxia (references below). Lesion or Cooling of Ventral Medullary Surface Areas
Both in the awake and anesthetized cat, lesioning of specific subretrofacial areas on the ventral medullary surface designated as area S and RetroTrapezoid Nucleus (RTN), respectively, virtually abolishes the ventilatory or phrenic nerve response to CO2 [175–179]. Studies using focal cooling of caudal, intermediate and more rostral ventral medullary surface areas showed profound effects on the pattern of breathing and CO2 sensitivity in cat (e.g., [180–182]), dog [183], and rabbit [184]. These studies were interpreted to mean that the central chemoreceptors were the main responsive elements in the CO2 response and also that at least one of the chemosensitive areas (area S) would serve as an integrator for all afferent chemosensory input [175,181]. More recent data obtained from awake goats after cooling the rostral M and the caudal M-rostral S areas could not confirm this hypothesis because the cooling did not eliminate CO2 sensitivity, even after carotid body denervation [185]. Also, in the same awake animal preparation, bilateral injection of an unselective excitatory amino acid receptor antagonist did not abolish the CO2 response but only reduced it by about 40% [186]. The latter findings are consistent with the view that if focal cooling and treatment of the classical chemosensitive areas with neurotoxins are not able to reduce the CO2 response by more than 60%, the remaining sensitivity can be explained by the presence of central chemoreceptors elsewhere. Thus local lesions or cooling are inappropriate instruments to estimate the overall contribution of central chemoreceptors to the CO2 response in normal conditions. In addition, following lesions (but also following carotid body denervation; see below), any possible peripheral-central interaction could disappear. However, the effects of focal cooling (and stimulation) have revealed some other
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interesting features about the role of ventral medullary chemosensitive areas, because these effects strongly depend on the arousal state of the animals (see below). Carotid Body Denervation
Shortly after Carotid Body Denervation (CBD) in awake adult rats [187], anesthetized and awake cats [188,189], awake dogs [190,191], and goats [192], but not in anesthetized rabbits [193], the ventilatory response to CO2 can be attenuated as much as 20–60%. The effects of CBD, however, are time- and age-dependent. In the goat, for example, CO2 sensitivity returns gradually (i.e., over about 15 days) to normal [192]. Over about three weeks, a similar recovery could not be observed in awake dogs showing a depression of the CO2 response by about 40%, two days after CBD [191]. This estimation, however, was made by using a modified Read rebreathing technique with different step sizes in CO2 and possibly different cerebral blood flow before and after CBD. This may lead to considerable differences between steady-state and rebreathing slopes [21]. After CBD in neonatal goats, piglets and rats, a normal CO2 sensitivity appears to have developed between three weeks and three months after the denervation [194]. These time- and age-dependent effects of CBD on the CO2 response make it questionable whether denervated animals (or carotid-body-resected humans) can provide a realistic picture of the normal contribution of the central chemoreceptors to total CO2 sensitivity. In addition, if the peripheral chemoreceptors provide a tonic facilitatory input to the central chemoreceptors, then an acute resetting by CBD of the central chemoreceptor gain cannot be ruled out and central CO2 sensitivity could be underestimated [195,196]. A tonic input of the carotid bodies into the RostroVentroLateral Medulla (RVLM) could be mediated by setting the local balance between glutamate and GABA turnover [197]. Note the many connections that have been found to exist between the caudal, commissural and medial subnuclei of the NTS on one hand and the RVLM, where many central chemoreceptors are located, on the other [84]. Note also that compared with normoxia, the number of noncatecholaminergic neurons activated by hypercapnia (inhalation of 15% CO2) in the RVLM is substantially reduced in hyperoxia, suggesting a reduced facilitatory input from the peripheral chemoreceptors in this condition [84]. If it can be ruled out that the recovery of the CO2 response after CBD is due to recovery of a peripheral chemoreflex originating in the aortic or carotid bodies, then the denervation experiments are an excellent means to show a remarkable CNS plasticity that may reside in the central chemosensitive network, the mechanism of which remains to be determined [192,194].
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Teppema and Dahan Selective Perfusion of Carotid Bodies and Brainstem
Selective perfusion of carotid bodies and brainstem is an elegant means to study separate central and peripheral contributions to total CO2 sensitivity. Studies with artificial brainstem perfusion in the anesthetized cat have revealed a contribution of the peripheral chemoreceptors as large as 20–58% (mean 48%) during hyperoxia [198]. In dogs undergoing selective perfusion of their carotid bodies, similar results were obtained [199]. Theoretically, a low stimulus level at the carotid bodies in these preparations could also diminish or remove a tonic facilitatory input to the central chemoreceptors and reduce central CO2 sensitivity. The pontomedullary perfusion experiments in the cat, however, have shown a constant central CO2 sensitivity over a wide range of peripheral CO2 tensions [198], suggesting the absence of a peripheral-central CO2 interaction. Dynamic End-tidal Forcing
This approach is based on the different speeds of response of the fast peripheral and slow central chemoreflex loops so that the responses of both reflex loops upon a step change in end-tidal PCO2 would be separable [23,24]. In both the anesthetized cat and humans (awake and sedated), the dynamic ventilatory response to square-wave changes in end-tidal CO2 fits a twocompartment model comprising a fast and slow compartment quite well [23,24]. In the anesthetized cat, the technique was validated by showing that the magnitudes of both components closely corresponded to those obtained during artificial brainstem perfusion [24]. The dynamic end-tidal forcing technique was refined by Pedersen et al. [200] by varying the end-tidal PCO2 with a multi-frequency binary sequence (MFBS), by which the identification and quantitative estimation of the fast peripheral component is improved. Generally all these dynamic studies yielded at least a 50–80% contribution of the central chemoreflex loop while that from the peripheral chemoreflex loop varied between 15 and 50%, with greater contribution at lower PO2 levels. In humans, the peripheral contribution to the total CO2 response during hyperoxia (estimated with MFBS and square-wave changes in endtidal PCO2, respectively) can be as large as 27% at a PO2 background of 200 Torr and 13% at a PO2 greater than 500 Torr [200,201]. B. Central Chemoreceptors in Metabolic Acid-Base Disturbances
The mechanisms by which the body defends against metabolic acid-base disturbances is a complex interplay between intra- and extracellular buffering, adjustment of acid excretion by the kidneys, and adjustment of CO2 removal by the lungs. Ventilatory adaptation to metabolic acid-base alterations is mediated via the peripheral and central chemoreceptors [202].
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The fact that the central chemoreceptors have an impressively high pH sensitivity does not mean that they are responsible for the ventilatory response to a metabolic acidosis or alkalosis. Using the arterial pH as an independent variable shows that the arterial chemoreceptors have a pH sensitivity twice as high as the central chemoreceptors [203]. The crucial factor here is the extent to which and how fast a primary metabolic alteration in arterial pH is reflected in the medullary extracellular fluid. Upon a bolus infusion of NaHCO3 or HCl performed in isocapnic conditions, this appears to be about 30% in the anesthetized cat [204,205], with a time course rapid enough to initiate fast ventilatory changes [205]. In a closed-loop situation, however, when the arterial PCO2 is not clamped as in the above cat experiments, metabolic arterial pH changes develop more slowly. In this situation, the peripheral chemoreceptors sense the change in pH, initiate a change in ventilation and induce a change in arterial PCO2 that will be rapidly followed by a change in medullary ECF PCO2. Thus in the acute phase of a metabolic acid-base disturbance, the medullary pHe (and pHi, but to a lesser degree) will undergo a paradoxical change and the central chemoreceptors will dampen down the ventilatory response initiated by the peripheral chemoreceptors that sense the original disturbance [203,206]. In the chronic phase, this dampening effect may gradually extinguish if compensatory changes in medullary extracellular bicarbonate concentration occur which would tend to normalize local extracellular pH. If a primary acidosis would exist in the brain, the situation would be more complex (cf., a condition of brain hypoxia where ventilatory output is reduced; see above). After CBD, the mechanism of ventilatory adaptation to a metabolic pH disturbance will be quite different, because in this case the central chemoreceptors will initiate a response depending on the extent to which the original disturbance penetrates into the CNS. C. Central Chemoreceptors and Tonic Drive to Breath
The classical areas L, S and M on the ventral medullary surface were originally described as regions with chemosensitive (M and L) and integrative (S) properties [7]. The most complex area of these is area S, which forms the ventral border of the RVLM with its known crucial role in the mediation of pain and the control of cardiovascular and arousal functions [84]. As discussed above, lesions or cooling of the classical areas (particularly S) have profound respiratory depressant effects in anesthetized cats, dogs and rabbits. In the anesthetized goat, focal bilateral cooling of area S and M for 30 s causes an apneic episode that outlasts the period of cooling [207]. In the anesthetized rat and cat, local lesions in the RTN and retrofacial area (which belong to the RVLM and partly overlap area S) cause a decrease in phrenic activity and CO2 sensitivity [177–179,208]. Cooling rostroventral (presumably M and S) chemosensitive areas in the cat
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releases a tonic activity in expiratory muscles [182]. Apart from chemosensitive cells, the RTN also contains neurons with respiratory-modulated activity [209,210]. These studies all indicate that the RVLM, particularly the superficial area extending from the rostral part of area S to the caudal region of area M (containing the RTN) are not only chemosensitive but also provide a tonic drive to the respiratory controller and even may influence the pattern of breathing. The total tonic input to the respiratory controller is composed of inputs from cortical and limbic structures, the reticular activating system, mechanoafferents from nonrespiratory muscles, and from the peripheral and central chemoreceptors. When during anesthesia and, to a lesser degree, during sleep the cortical, limbic and mechanoafferent inputs are reduced, the relative importance of chemical feedback from the chemoreceptors will increase, which is consistent with the current idea that during (NREM) sleep and anesthesia ventilation is predominantly under metabolic control. The dependency of ventilation on central chemoreceptor drive in sleep and anesthesia appears from many animal data. For example, while focal cooling of the rostral chemosensitive areas of goats resulted in complete apnea during anesthesia, during wakefulness a ventilatory depression of only 30% was induced. This effect tended to be somewhat greater during NREM sleep [185,207,211,212]. The crucial importance of central chemoreceptor drive during sleep is also illustrated in patients suffering from the congenital central hypoventilation syndrome (CCHS). Generally, these patients do not show appreciable sensitivity to CO2 during wakefulness and sleep. However, when they are aroused, exercise or execute cognitive tasks, most of them breathe adequately and maintain relatively normal blood gases [213]. When they are asleep, however, they may severely hypoventilate, resulting in high PCO2 levels [213,214; see also Chapter 9 by Gozal, this volume] (note that some CCHS patients also hypoventilate when awake; [214]). Hypercapniainduced arousal is another interesting phenomenon in CCHS patients lacking ventilatory CO2 sensitivity [215]. Could this be mediated by a subset of the many neurons in the brainstem activated during hypercapnia? Note that the picture of these neurons in the brain activated by CO2 is incomplete. Recent fMRI data indicate that hypercapnic challenges elicit discrete changes in activity in multiple brain sites including diencephalic and (sub)cortical structures [216]. Many of these regions are also activated in CCHS patients, but some pontomedullary, cerebellar and (sub)cortical regions showed a reduced hypercapnia-induced activation [217]. More illustrative evidence for the importance of central chemical drive during sleep (or the importance of wakefulness drive when awake) comes from patients with unilateral focal lesions in the rostrolateral medulla. During wakefulness, most of the nine patients studied by Morell et al. [218] maintained a normal PCO2 during rest and exercise, but had a lower than
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normal CO2 sensitivity. All patients, however, displayed a disturbed and fragmented sleep pattern, often accompanied by obstructive apneas [218,219]. D. Central Chemosensitive Areas Throughout the Brainstem: Different Functions for Distinct Areas?
An intriguing property of the central chemoreceptors is their widespread distribution throughout the brainstem. It was suggested that this distribution may have evolutionary grounds based on gradual development of functions associated with the transition from water- to air-breathing, temperature regulation and sleep [124]. That so many neurons are activated by hypercapnia (data from Fos studies) is not very surprising, because apart from respiratory responses, CO2 also elicits cardiovascular, endocrine and thermoregulatory reactions. Consequently, not all brainstem neurons that stain for Fos upon hypercapnic exposure will be respiratory CO 2 chemoreceptors. On the other hand, it is relevant to consider that at least in the medulla, all chemosensitive areas described with the aid of focal stimulations lie well within the regions containing Fos-positive neurons. Suppose then that all these distinct areas serve to influence respiratory output during hypercapnia (which, given the invariable respiratory effects of focal lesions and stimulations seems a reasonable assumption, at least in anesthetized animals), what would be the advantage of their distant locations? It is evident that by this organization various specific respiratory adaptations can be effectuated simultaneously, because with their own efferent connections, all areas involved have distinct specific functions in controlling the extremely complex breathing apparatus. Of utmost importance is also the fact that all the chemosensitive areas have different afferent connections, some of which are clearly under the influence of the arousal state [84]. This then could have the consequence that some areas behave differently in different arousal states, and indeed this appears to be the case. A first example is the RVLM. The different effects of superficial RVLM cooling in the goat during anesthesia and wakefulness may be caused by a decreased tonic input of chemosensitive neurons to the respiratory controller during wakefulness. This could be due to inhibitory influences from the cortex on RTN neurons, which of course does not preclude a simultaneous powerful facilitatory influence of the cortex on respiratory neurons. It is also interesting to speculate about a possible contribution to the observed phenomena of the locus coeruleus with its extensive noradrenergic projections throughout the brainstem [220]. Locus Coeruleus (LC) neurons are activated during hypercapnia in awake but not anesthetized rats [84,96]. Do noradrenergic projections from the (chemosensitive) LC influence the excitability of RVLM neurons, and if yes, in
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NTS −11.00 VII
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Figure 2.7 CO2 microdialysis in the retrotrapezoid nucleus (RTN, left panel) and ventral medullary raphe (right panel) in awake and sleeping rats. CO2 has stimulatory effects (increase in tidal volume) in the RTN during wakefulness but not sleep. In raphe the opposite occurs: stimulation (increase in breathing frequency) during sleep but not while awake. NTS: nucleus tractus solitarius; VII: motor nucleus of the facial nerve (Data from Ref. 227).
what sense? (For a review of central noradrenergic effects on breathing, see [221].) An alternative explanation for the above findings in goats could be a smaller relative contribution of area S-M to total central chemical drive during wakefulness, so that removal of a given amount of their output would have a smaller effect on ventilation than during anesthesia or sleep. A surprising finding in rats (in the light of the above results of cooling in goats) was that microdialysis of CO2 in the RTN increased breathing in awake but not sleeping animals (Figure 2.7, left panel; [63,64]). In anesthetized animals, however, focal CO2 in the RTN increased phrenic activity [61]. At first sight these results in rats and goats seem hard to reconcile, but they may not necessarily prove to be conflicting. Some crucial questions to be answered here are: 1) What is the effect of sleep vs. anesthesia on the CO2 sensitivity of RTN neurons? 2) Do sleep and anesthesia alter the sensitivity of respiratory neurons to input from the RTN
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and other chemosensitive regions? 3) What are the effects of sleep and anesthesia further downstream in the ventilatory control system? The stimulatory effect of focal CO2 in the RTN in awake but not sleeping animals could offer one of the possible explanations for a generally observed higher CO2 sensitivity during wakefulness when compared with sleep. Another example of a chemosensitive area with a possibly specific role in respiratory regulation is the ventral medullary raphe region with a high concentration of serotonergic chemosensitive neurons. In the rat, microdialysis of CO2 in this region increases breathing frequency in sleep but not wakefulness (Figure 2.7, right panel; [62]). During NREM sleep, an increased GABA-ergic input into serotonergic raphe neurons may be responsible for their reduced activity in this state [222,223]. Assuming that this GABA-ergic input leads to hyperpolarization, this may lead to a situation in which the conductance of ion channels in the membrane of these neurons is changed (hyperpolarization, apart from probably direct effect of GABA on specific ionic conductances). If CO2/Hþ would act on raphe neurons by a direct effect on voltage-dependent ion channels that open when the membrane hyperpolarizes, this could lead to a situation in which a CO2/ Hþ-induced effect either could get lost or rather arise following a membrane hyperpolarization. Kir channels could be a candidate for such channels because they open with hyperpolarization [37], may be very pH sensitive [168], and may close upon a decrease in pH which would restore the membrane potential to the level existing without the GABA-ergic input. Various facilitatory inputs that impinge on raphe neurons during wakefulness could be filtered away during sleep, thus allowing CO2 to increase their output. Theoretically, for raphe neurons it seems to make sense to increase their output upon a rise in PCO2 during sleep. By providing a facilitatory input to hypoglossal motoneurons, serotonergic raphe neurons play a role in regulating upper airway patency [224]. During sleep, a decrease in activity of these cells may eventualy result in reduced genioglossal activity [225]. In man, upper airway mechanoreceptors are much less active in NREM sleep than when awake, with a low genioglossal response to large inspiratory resistive loads [226]. By increasing their CO2 sensitivity during sleep, raphe neurons could thus help to prevent, reduce or reverse obstruction of the upper airways. By increasing respiratory frequency in response to CO2 during sleep [62,227], raphe neurons could also reduce the negative airway pressure per breath that is needed to achieve a given level of alveolar ventilation.
References 1.
Leusen, I.R., Chemosensitivity of the respiratory centre: Influence of CO2 in the cerebral ventricles on respiration, Am. J. Physiol. 176, 39–44, 1954.
56
Teppema and Dahan
2.
Pappenheimer, J.R., Fencl, V., Heisy, S.R. and Held, D., Role of cerebral fluids in the control of respiration as studied in unanesthetized goats, Am. J. Physiol. 208, 436–450, 1965. Fencl, V., Miller, T.B. and Pappenheimer, J.R., Studies on the respiratory response to disturbance of acid-base balance, with deductions concerning the ionic composition of cerebral interstitial fluid, Am. J. Physiol. 210, 459–472, 1966. Berndt, J., Berger, W., Berger, K. and Schmiedt, M., Untersuchungen zum zentralen chemosensibelen Mechanismus der Atmung, II, Die Steuerung der Atmung durch das extracellula¨re pH im Gewebe der Medulla Oblongata, Pflu¨gers Arch. 332, 146–170, 1972. Loeschcke, H.H., The respiratory control system: Analysis of steady-state solutions for metabolic and respiratory acidosis-alkalosis and increased metabolism, Pflu¨gers Arch. 341, 23–42, 1973. Berkenbosch, A., DeGoede, J., Olievier, C.N., Quanjer, P.H., Perk, J.H., Philips, L. and Rancuret, M.M., Influence of CSF bicarbonate concentration on the ventilatory response to CO2 in relation to the location of the central chemoreceptors, Respir. Physiol. 35, 215–236, 1978. Schlaefke, M.E., Central chemosensitivity: A respiratory drive, Rev. Physiol. Biochem. Pharmac. 90, 171–244, 1981. Loeschcke, H.H., Central CO2 sensitivity and the reaction theory (Review), J. Physiol. (Lond.) 332, 1–24, 1982. Eldridge, F., Kiley, J.P. and Millhorn, D.E., Responses to medullary hydrogen ion changes in cats: Different effects of respiratory and metabolic acidoses, J. Physiol. (Lond.) 358, 285–297, 1985. Teppema, L.J., Barts, P.W.J.A., Folgering, H.T. and Evers, J.A.M., Effects of respiratory and (isocapnic) metabolic arterial acid-base disturbances on medullary extracellular fluid pH and ventilation in cats, Respir. Physiol. 53, 379–395, 1983. Shams, H., Differential effects of CO2 and Hþas central stimuli of respiration in the cat, J. Appl. Physiol. 58, 357–364, 1985. Marino, P.L. and Lamb, T.W., Effects of CO2 and extracellular Hþ iontophoresis on single cell activity in the cat brainstem, J. Appl. Physiol. 38, 688–695, 1975. Kawai, A., Ballantyne, D., Mu¨ckenhoff, K. and Scheid, P., Chemosensitive medullary neurons in the brainstem-spinal cord preparation of the neonatal rat, J. Physiol. 492, 277–292, 1996. Richerson, G.B., Response to CO2 of neurons in the rostral ventral medulla in vitro, J. Neurophysiol. 73, 933–944, 1995. Richerson, G.B., Wang, G., Tiwari, J. and Bradley, S.R., Chemosensitivity of serotonergic neurons in the rostral ventral medulla, Respir. Physiol. 129, 175–191, 2001. Okada, Y., Chen, Z., Jiang, W., Kuwana, S. and Eldridge, F.L., Anatomical arrangement of hypercapnia-activated cells in the superficial ventral medulla of rats, J. Appl. Physiol. 93, 427–439, 2002. Gorcs, T.J., Liposits, Z., Palay, S.L. and Chan-Palay, V., Serotonin neurons on the ventral brain surface, Proc. Natl. Acad. Sci. USA 82, 7449–7452, 1985.
3.
4.
5.
6.
7. 8. 9.
10.
11. 12.
13.
14. 15.
16.
17.
Central Chemoreceptors 18.
19. 20. 21.
22.
23. 24.
25.
26.
27.
28.
29.
30.
31. 32.
33.
34.
57
Bradley, S.R., Pieribone, V.A., Wang, W., Severson, C.A., Jacob, R.A. and Richerson, G.B., Chemosensitive serotonergic neurons closely associated with large medullary arteries, Nat. Neurosci. 5, 401–402, 2002. Read, D.J.C., A clinical method for assessing the ventilatory response to carbon dioxide, Australas. Ann. Med. 16, 20–32, 1967. Read, D.J.C. and Leigh, J., Blood-brain tissue PCO2 relationships and ventilation during rebreathing, J. Appl. Physiol. 23, 53–70, 1967. Berkenbosch, A., Bovil, J.G., Dahan. A., DeGoede, J. and Olievier, I.C.W., Ventilatory sensitivities from Read’s rebreathing method and the steady-state method are not equal, J. Physiol. (Lond.) 411, 367–377, 1989. Dutton, R.E., Davies, D.G., Ghatak, P.K. and Fitzgerald, R.S., Respiration during transient perfusion of vertebral arteries with hypocapnic blood, Am. J. Physiol. 217, 1178–1182, 1969. Swanson, G.D. and Bellville, J.W., Step changes in end-tidal CO2: Methods and implications, J. Appl. Physiol. 39, 377–385, 1975. DeGoede, J., Berkenbosch, A., Ward, D.S., Bellville, J.W. and Olievier, C.N., Comparison of chemoreflex gains obtained with two different methods in cats, J. Appl. Physiol. 59, 170–179, 1985. Teppema, L.J., Vis, A., Evers, J.A.M. and Folgering, H.T., Dynamics of brain extracellular fluid pH and phrenic nerve activity in cats after end-tidal CO2 forcing, Respir. Physiol. 50, 359–380, 1982. Berkenbosch, A., Olievier, C.N., DeGoede, J. and Kruyt, E.W., Effect on ventilation of papaverine administered to the brainstem of the anaesthetized cat, J. Physiol. (Lond.) 443, 457–468, 1991. VanBeek, J.H.G.M., Berkenbosch, A., DeGoede, J. and Olievier, C.N., Effects of brainstem hypoxaemia on the regulation of breathing, Respir. Physiol. 57, 171–188, 1984. Teppema, L.J., Rochette, F. and Demedts, M., Ventilatory response to carbonic anhydrase inhibition in cats: effects of acetazolamide in intact vs. peripherally chemodenervated animals, Respir. Physiol. 74, 373–382, 1988. Teppema, L.J., Rochette, F. and Demedts, M., Effects of acetazolamide on medullary extracellular pH and PCO2 and on ventilation in peripherally chemodenervated cats, Pflu¨gers. Arch. 415, 519–525, 1990. Teppema, L., Berkenbosch, A., DeGoede, J. and Olievier, C., Carbonic anhydrase and the control of breathing: Different effects of benzolamide and methazolamide in the anesthetized cat, J. Physiol. 488, 767–777, 1995. Cragg, P., Patterson, L. and Purves, M.J., The pH of brain extracellular fluid in the cat, J. Physiol. (Lond.) 272, 137–166, 1977. Oyamada, Y., Andrzejewski, M., Mu¨ckenhof, K., Scheid, P. and Ballantyne, D., Locus coeruleus neurons in vitro: pH sensitive oscillations of membrane potential in an electrically coupled network, Respir. Physiol. 118, 131–147, 1999. Andrzejewski, M., Mu¨ckenhof, K., Scheid, P. and Ballantyne, D., Synchronized rhythms in chemosensitive neurons of the locus coeruleus in the absence of chemical synaptic transmission, Respir. Physiol. 129, 123–140, 2001. Dean, J.B., Bayliss, D.A., Erickson, J.T., Lawing, W.L. and Millhorn, D.E., Depolarization and stimulation of neurons in nucleus tractus solitarii by carbon
58
35.
36. 37.
38. 39.
40. 41. 42. 43.
44.
45. 46. 47. 48. 49.
50.
51. 52.
Teppema and Dahan dioxide does not require chemical synaptic input, Neuroscience 36, 207–216, 1990. Rigatto, H., Fitzgerald, S.C., Willis, M.A. and Yu, C., In search of the central repiratory neurons: II, Electrophysiologic studies of medullary fetal cells inherently sensitive to CO2 and low pH, J. Neurosci. Res. 33, 590–597, 1992. Dillon, G.H. and Waldrop. T.G., In vitro responses of caudal hypothalamic neurons to hypoxia and hypercapnia, Neuroscience 51, 941–950, 1992. Pineda, J. and Aghajanian, G.K., Carbon dioxide regulates the tonic activity of locus coeruleus neurons by modulating a proton- and polyamine-sensitive inward rectifying potassium current, Neuroscience 77, 723–743, 1997. Wang, W., Pizzonia, J.H. and Richerson, G.B., Chemosensitivity of rat medullary raphe neurons in primary tissue culture, J. Physiol. (Lond.) 511, 433–450, 1998. Wang, W., Tiwari, J.K., Bradley, S.R., Zaykin, A.V. and Richerson, G.B., Acidosis-stimulated neurons of the medullary raphe are serotonergic, J. Neurophysiol. 85, 2224–2235, 2001. Ridderstra˚le, Y. and Hanson, M.A., Histochemical localization of carbonic anhydrase in the carotid body, Proc. N.Y. Acad. Sci. 429, 398–400, 1984. Rigual, C., Iniguez, C., Carreres, J. and Gonzalez, C., Carbonic anhydrase in the carotid body and sinus nerve, Histochem. 82, 577–580, 1985. Nurse, C.A., Carbonic anhydrase and neuronal enzymes in cultured glomus cells of the carotid body of the rat, Cell. Tiss. Res. 261, 65–71, 1990. Botre, F., Botre, C., Greco, A., Data, P.G., Di Giulio, C. and Morelli, L., Potentiometric determination of carbonic anhydrase activity in rabbit carotid arteries: Comparison among normoxic, hyperoxic and hypoxic animals, Neurosci. Lett. 166, 126–130, 1994. Coates, E.L., Li, A. and Nattie, E.E., Acetazolamide on the ventral medulla of the cat increases phrenic output and delays the ventilatory response to CO2, J. Physiol. 441, 433–451, 1991. Coates, E.L., Li, A. and Nattie, E.E., Widespread sites of brain stem ventilatory chemoreceptors, J. Appl. Physiol. 75, 5–14, 1993. Bernard, D.G., Li, A. and Nattie, E.E., Evidence for central chemoreception in midline raphe, J. Appl. Physiol. 80, 108–115, 1996. Nattie, E.E. and Li, A., Central chemoreception in the region of the ventral respiratory group in the rat, J. Appl. Physiol. 81, 1987–1995, 1996. Solomon, I.C., Edelman, N.H. and O’Neal, M.H., CO2/Hþ chemoreception in the cat pre-Bo¨tzinger complex in vivo, J. Appl. Physiol. 88, 1996–2007, 2000. Xu, F., Zhang, Z. and Frazier, D.T., Microinjection of acetazolamide into the fastigial nucleus augments respiratory output in the rat, J. Appl. Physiol. 91, 2342–2350, 2001. Wang, W., Bradley, S.R. and Richerson, G.B., Quantification of the response of rat medullary raphe neurons to independent changes in pHo and PCO2, J. Physiol. (Lond.) 540, 951–970, 2002. Ridderstra˚le, Y. and Wistrand, P.J., Membrane-associated carbonic anhydrase activity in the brain of CA II-deficient mice, J. Neurocytol. 29, 263–269, 2000. Smith, J.C., Ellenberger, H., Ballanyi, K., Richter, D. and Feldman, J., Pre-Bo¨tzinger complex: A brainstem region that may generate respiratory rhythm in mammals, Science 254, 726–729, 1991.
Central Chemoreceptors 53.
54.
55. 56.
57.
58.
59.
60.
61. 62. 63.
64.
65.
66.
67.
68.
59
Buckler, K.J., Vaughan-Jones, R.D., Peers, C. and Nye, P.C., Intracellular pH and its regulation in type I carotid body cells of the neonatal rat, J. Physiol. 436, 107–129, 1991. Hanson, M.A., Nye, P.C. and Torrance, R.W., The exodus of an extracellular bicarbonate theory of chemoreception and the genesis of an intracellular one, in Arterial Chemoreceptors, Belmonte, C., Pallot, D., Acker, H. and Fidone, S., eds., Leicester, Leicester University Press, pp. 403–416, 1981. Itturiaga, R., Lahiri, S. and Mokashi, A., Carbonic anhydrase and chemoreception in the carotid body, Am. J. Physiol. 265, C565–C573, 1991. Lahiri, S., Itturiaga, R., Mokashi, A., Botre, F., Chugh, D. and Osanai, S., Adaptation to hypercapnia vs. intracellular pH in cat carotid body: Responses in vitro, J. Appl. Physiol. 80, 1090–1099, 1996. Adams, J.M. and Johnson, N.L., Inhibiting carbonic anhydrase in brain tissue increases the respiratory response to rebreathing CO2, Brain Res. 519, 23–28, 1990. Pickkers, P., Garcha R.S., Schachter, M., Smits, P. and Hughes, A.D., Inhibition of carbonic anhydrase accounts for the direct vascular effects of hydrochlorothiazide. Hypertension 33, 1043-1048, 1999. Pickkers, P., Hughes, A.D., Russel, F.G.M., Thien, Th. and Smits, P., In vivo evidence for KCa channel opening properties of acetazolamide in the human vasculature, Br. J. Pharmacol. 132, 443–450, 2001. Teppema, L.J., Dahan, A. and Olievier, C.N., Low-dose acetazolamide reduces CO2-O2 stimulus interaction within the peripheral chemoreceptors in the anaesthetised cat, J. Physiol. (Lond.) 537, 221–229, 2001. Li, A. and Nattie, E.E., Focal central chemoreceptor sensitivity in the RTN studied with a CO2 diffusion pipette in vivo, J. Appl. Physiol. 83, 420–428, 1997. Nattie, E.E. and Li, A., CO2 dialysis in the medullary raphe of the rat increases ventilation in sleep, J. Appl. Physiol. 90, 1247–1257, 2001. Li, A., Randall, M. and Nattie, E.E., CO2 microdialysis in retrotrapezoid nucleus of the rat increases breathing in wakefulness but not in sleep, J. Appl. Physiol. 87, 910–919, 1999. Li, A. and Nattie, E., CO2 dialysis in one chemoreceptor site, the RTN: Stimulus intensity and sensitivity in the awake rat, Respir. Physiol. Neurobiol. 133, 11–22, 2002. Nattie, E. and Li, A., CO2 dialysis in nucleus tractus solitarius region of rat increases ventilation in sleep and wakefulness, J. Appl. Physiol. 92, 2119–2130, 2002. Ridderstra˚le, Y. and Wistrand, P.J., Carbonic anhydrase isoforms in the mammalian nervous system, in pH and Brain Function, Kaila, K. and Ransom, B.R., eds., New York, John Wiley Liss, pp. 21–43, 1998. Agnati, L.F., Tinner, B., Staines, W.A., Va¨a¨na¨nen, K. and Fuxe, K., On the cellular localization and distribution of carbonic anhydrase II immunoreactivity in the rat brain, Brain Res. 676, 10–24, 1995. Ghandour, M.S., Vincendon, G., Gombos, G., Limozin, N., Filippi, D., Dalmasso, C. and Laurent, G., Carbonic anhydrase and oligodendroglia in developing rat cerebellum: A biochemical and immunohistochemical study, Dev. Biol. 77, 73–83, 1980.
60
Teppema and Dahan
69.
Ghandour, M.S., Langley, O.K., Zhu, X.L., Waheed, A. and Sly, W.S., Carbonic anhydrase IV on brain capillary endothelial cells: A marker associated with the blood-brain barrier, Proc. Nat. Acad. Sci. USA, 89, 6823–6827, 1992. Brion, L.P., Suarez, C., Zhang, H. and Cammer, W., Up-regulation of carbonic anhydrase iso-enzyme IV in CNS myelin in mice genetically deficient in carbonic anhydrase II, J. Neurocytochem. 63, 360–366, 1994. Cammer, W., Zhang, H. and Tansey, F.A., Effects of carbonic anhydrase II (CAII) deficiency on CNS structure and function in myelin-deficient CAIIdeficient double mutant mice, J. Neurosci. Res. 40, 451–457, 1995. Cammer, W.B. and Brion, L.P., Carbonic anhydrase in the nervous system, in The Carbonic Anhydrases, Chegwidden, W.R., Carter, N.D. and Edwards, Y.H., eds., New Horizons, Basel, Birha¨user Verlag, pp. 475–489, 2000. Ridderstra˚le, Y. and Hanson, M.A., Histochemical study of the distribution of carbonic anhydrase in the cat brain, Act. Physiol. Scand. 124, 557–564, 1985. Neubauer, J.A., Carbonic anhydrase and sensory function in the central nervous system, in The Carbonic Anhydrases, Dodgson, S., ed., New York, Plenum, pp. 319–323, 1991. Hanley, M.R., Proto-oncogenes in the nervous system, Neuron 1, 175–182, 1988. Morgan, J.I. and Curran, T., Stimulus-transcription coupling in neurons: Role of cellular immediate-early genes, TINS 12, 459–462, 1989. Morgan, J.I. and Curran, T., Stimulus-transcription coupling in the nervous system: involvement of the inducible proto-oncogenes fos and jun, Annu. Rev. Neurosci. 14, 421–451, 1991. Morgan, J.I. and Curran, T., Role of ion flux in the control of c-fos expression, Nature 322, 552–555, 1986. Sheng, M. and Greenberg, M.E., The regulation and function of c-fos and other immediate early genes in the nervous system, Neuron 4, 477–485, 1990. Sato, M., Severinghaus, J.W. and Basbaum, A.I., Medullary CO2 receptor neuron identification by c-fos immunocytochemistry, J. Appl. Physiol. 73, 96– 100, 1992. Swanson, L.W., Brain maps, in Structure of the Rat Brain, New York, Elsevier, 1992. Teppema, L.J., Berkenbosch, A., Veening, J.G. and Olievier, C.N., Hypercapnia induces c-fos expression in neurons of retrotrapezoid nucleus in cats, Brain Res. 635, 353–355, 1994. Teppema, L.J., Veening, J.G. and Berkenbosch, A., Expression of c-fos in the brain stem of rats during hypercapnia, Adv. Exp. Med. Biol. 393, 47–51, 1995. Teppema, L.J., Veening, J.G., Kranenburg, A., Dahan, A., Berkenbosch, A. and Olievier, C., Expression of c-fos in the rat brainstem after exposure to hypoxia and to normoxic and hyperoxic hypercapnia, J. Comp. Neurol. 388, 169–190, 1997. Teppema, L.J., Kranenburg, A., Veening. J.G. and Berkenbosch. A., Expression of c-fos in hypothalamus and forebrain of rats during hypoxia and hypercapnia, Proc. XIIIth International Symposium on Arterial Chemoreceptors, Santiago, Chile, 78, 1996.
70.
71.
72.
73. 74.
75. 76. 77.
78. 79. 80.
81. 82.
83.
84.
85.
Central Chemoreceptors 86.
87.
88.
89.
90.
91.
92.
93.
94. 95. 96.
97.
98.
99.
100. 101.
61
Jansen, A.H., Liu, P., Weisman, H., Chernick, V. and Nance, D.M., Effect of sinus denervation and vagotomy on c-fos expression in the nucleus tractus solitarius after exposure to CO2, Pflu¨gers Archiv. 431, 876–881, 1996. Haxhiu, M.A., Yung, K., Erokwu. B and Cherniack, N.S., CO2 induced c-fos expression in CNS catecholaminergic neurons, Respir. Physiol. 105, 35–45, 1996. Berquin, P., Bodineau, L., Gros, F. and Larnicol, N., Brainstem and hypothalamic areas involved in respiratory chemoreflexes: A Fos study in adult rats, Brain Res. 857, 30–40, 2000. Larnicol, N., Wallois, F., Berquin, P., Gros, F. and Rose D., c-fos-like immunoreactivity in the cat’s neuraxis following moderate hypoxia or hypercapnia, J. Physiol. (Paris) 88, 81–88, 1994. Miura, M., Okada, J., Takayama, K. and Suzuki, T., Neuronal expression of Fos and Jun protein in the rat medulla and spinal cord after anoxic and hypercapnic stimulations, Neurosci. Lett. 178, 227–230, 1994. Miura, M., Okada, J. and Kanazawa, M., Topology and immunohistochemistry of proto-sensitive neurons in the ventral medullary surface of rats, Brain Res. 780, 34–45, 1998. Xu, F., Zhou, T., Gibson, T. and Frazier, D.T., Fastigial-nucleus mediated respiratory responses depend on the medullary gigantocellular nucleus, J. Appl. Physiol. 91, 1713–1722, 2001. Takayama, K., Suzuki, T. and Miura, M., The comparison of effects of various anesthetics on expression of Fos protein in the rat brain, Neurosci. Lett. 176, 59–62, 1994. Dragunow, M. and Faul, R., The use of c-fos as a metabolic marker in neuronal pathway tracing, J. Neurosci. Meth. 29, 261–265, 1989. Krukoff, T.L., Expression of c-fos in studies of central autonomic and sensory systems, Mol. Neurobiol. 7, 247–263, 1994. Teppema, L.J., Kranenburg, A. and Veening, J.G., Fos expression in hypothalamus and forebrain of conscious and anesthetized rats by exposure to hypercapnic air, J. FASEB 12(Pt 1 Suppl): 291, 1998. Erickson, J.T. and Millhorn, D.E., Hypoxia and electrical stimulation of the carotid sinus nerve induce fos-like immunoreactivity within catecholaminergic and serotonergic neurons of the rat brain stem, J. Comp. Neurol. 348, 161–182, 1994. Hoffman, G., Smith, M.S. and Verbalis, J.G., c-Fos and related immediate early gene products as markers of activity in neuroendocrine systems, Front. Neuroendocrinol. 14, 173–213, 1993. Douglas, R.M., Trouth, C.O., James, D., Sexcius, L.M., Kc, P., Dehkordi, O., Valladares, E.R. and Mckenzie, J.C., Decreased CSF pH at ventral brain stem induces widespread c-Fos immunoreactivity in rat brain neurons, J. Appl. Physiol. 90, 475–485, 2001. Kiwull-Scho¨ne, H. and Kiwull, P., Hypoxia and ‘‘the reaction theory’’ of central respiratory chemosensitivity, Adv. Exp. Med. Biol. 316, 347–357, 1992. Xu, F.D., Sato, M., Spellman, M.J., Mitchell, R.A. and Severinghaus, J.W., Topography of cat medullary surface hypoxic acidification, J. Appl. Physiol. 73, 2631–2637, 1992.
62
Teppema and Dahan
102. Xu, F.D., Spellman, M.J., Sato, M., Baumgartner, J.E., Ciricillo, S.F. and Severinghaus, J.W., Anomalous acidification of medullary ventral surface, J. Appl. Physiol. 71, 2211–2217, 1991. 103. Wiemann, M. and Bingmann, D., Ventrolateral neurons of medullary organotypic cultures: Intracellular pH regulation and bioelectric activity, Respir. Physiol. 129, 57–70, 2001. 104. Wiemann, M., Baker, R.E., Bonnet, U. and Bingmann, D., CO2 sensitive medullary neurons: Activation by intracellular activation, Neuroreport 9, 167–170, 1998. 105. Ritucci, N.A., Dean, J.B. and Putnam, R.W., Intracellular pH response to hypercapnia in neurons from chemosensitive areas of the medulla, Am. J. Physiol. 273, R433–R441, 1997. 106. Ritucci, N.A., Chambers-Kersh, L., Dean, J.B. and Putnam, R.W., Intracellular pH regulation in neurons from chemosensitive and nonchemosensitive areas of the medulla, Am. J. Physiol. 275, R1152–R1163, 1998. 107. Lyall, V., Feldman, G.M., Heck, G.L. and DeSimone, J.A., Effects of extracellular pH, PCO2 and HCO 3 on intracellular pH in isolated rat taste buds, Am. J. Physiol. 273, C1008–C1019, 1997. 108. Putnam, R.W., Intracellular pH regulation of neurons in chemosensitive and nonchemosensitive areas of brain slices, Respir. Physiol. 129, 37–56, 2001. 109. Ma, E. and Haddad, G.G., Expression and localization of Naþ/Hþ in rat central nervous system, Neuroscience 79, 591–601, 1997. 110. Kiwull-Scho¨ne, H., Wiemann, M., Frede, St., Bingmann, D., Wirth, K.J., Heinelt, U., Lang, H-J. and Kiwull, P., A novel inhibitor of the Naþ/Hþ exchanger type 3 activates the central respiratory CO2 response and lowers the apneic threshold. Am. J. Resp. Crit. Care Med. 164, 1303–1311, 2001. 111. Nottingham, S., Leiter, J.C., Wages, P., Buhay, S. and Erlichmann, J.S., Developmental changes in intracellular pH regulation in medullary neurons of the rat, Am. J. Physiol. 281: R1940–R1951, 2001. 112. Thomas, T., Ralevic, V., Gadd, C.A. and Spyer, K.M., Central chemoreception: a mechanism involving P2 purinoceptors localized in the ventrolateral medulla of the anaesthetetized rat, J. Physiol. (Lond.) 517, 899–905, 1999. 113. Thomas, T. and Spyer, K.M., ATP as a mediator of mammalian central chemoreception, J. Physiol. (Lond.) 532, 441–447, 2000. 114. Burton, M.D. and Kazemi, H., Neurotransmitters in central respiratory control, Respir. Physiol. 122, 111–121, 2000. 115. Burton, M.D., Nouri, K., Baichoo, S., Samuels-Toyloy, N. and Kazemi, H., Ventilatory output and acetylcholine: Perturbations in release and muscarinic receptor activation, J. Appl. Physiol. 77, 2275–2284, 1994. 116. Dev, N.B. and Loeschcke, H.H., Cholinergic mechanism involved in respiratory chemosensitivity in the cat., Pflu¨gers Arch. 379, 29–36, 1979. 117. Nattie, E.E., Wood, J., Mega, A. and Goritski, W., Rostral ventrolateral medulla muscarinic receptor involvement in central ventilatory chemosensitivity, J. Appl. Physiol. 66, 1462–1470, 1989. 118. Nattie, E.E. and Li, A., Ventral medulla site of muscarinic subtypes involved in cardiorespiratory control, J. Appl. Physiol. 69, 33–41, 1990.
Central Chemoreceptors
63
119. Nattie, E.E. and Li, A., Muscimol dialysis in the retrotrapezoid nucleus region inhibits breathing in the awake rat, J. Appl. Physiol. 89, 153–162, 2000. 120. Curran, A.K., Darnall, R.A., Filiano, J.J., Li, A. and Nattie, E.E., Muscimol dialysis in the rostral medulla reduces the ventilatory response to CO2 in awake and sleeping piglets, J. Appl. Physiol. 90, 971–980, 2001. 121. Fukuda, Y. and Loeschcke, H.H., Cholinergic mechanism involved in neuronal excitation by Hþ in rats in vitro, Pflu¨gers Arch. 379, 125–135, 1979. 122. Burton, M.D., Johnson, D.C. and Kazemi, H., CSF acidosis augments ventilation through cholinergic mechanisms. J. Appl. Physiol. 66, 2565–2572, 1989. 123. Nattie, E.E., Diethyl pyrocarbonate (an imidazole binding substance) inhibits rostral VLM CO2 sensitivity, J. Appl. Physiol. 64, 1600–1609, 1988. 124. Nattie, E.E., CO2 brainstem chemoreceptors and breathing, Progr. Neurobiol. 59, 299–331, 1999. 125. Gibson, J.S., Cossins, A.R. and Ellorey, J.C., Oxygen-sensitive membrane transporters in vertebrate red cells, J. Exp. Biol. 203, 1395–1407, 2000. 126. Yao, H., Gu, X.Q. and Haddad, G.G., The role of HCO 3 -dependent mechanisms on pHi regulation during O2 deprivation, Neuroscience 117, 29–35, 2003. 127. Deitmer, J.W., Strategies for metabolic exchange between glial cells and neurons, Respir. Physiol. 129, 71–81, 2001. 128. Dermietzel, R. and Spray, D.C., Gap junctions in the brain: where, what type, how many and why? Trends Neurosci. 16, 186–192, 1993. 129. Solomon, I.C., Halat, T.J., El-Maghrabi, M.R. and O’Neal MH III. Localization of connexin26 and connexin32 in putative CO2-sensitive brainstem regions in rat, Respir. Physiol. 129, 101–129, 2001. 130. Solomon, I.C., Halat, T.J., El-Maghrabi, M.R. and O’Neal MH III, Differential expression of connexin26 and connexin32 in the pre-Bo¨tzinger complex, J. Comp. Neurol. 440, 12–19, 2001. 131. Dean, J.B., Kinkade, E.A. and Putnam, R.W., Cell-cell coupling in CO2/Hþexcited neurons in brainstem slices, Respir. Physiol. 129, 83–100, 2001. 132. Huang, R.Q., Erlichman, J.S. and Dean, J.B., Cell-cell coupling between CO2 excited neurons in the dorsal medulla oblongata, Neuroscience 80, 41–57, 1997. 133. Peracchia, C., Sotkis, A., Wang, X.G., Peracchia, L.L. and Persechini, A., Calmoduline directly gates gap junction channels, J. Biol. Chem. 275, 26220–26224, 2000. 134. Spray, D.C., Stern, J.H., Harris, A.L. and Bennett, M.V.L., Gap junctional conductance: comparison of sensitivities to Hþ and Ca2þ ions, Proc. Natl. Acad. Sci. USA 79, 441–445, 1982. 135. Peracchia, C., Wang, X.G. and Peracchia, L.L., Is the chemical gate of connexins voltage sensitive? Behavior of Cx32 wild-type and mutant channels, Am. J. Physiol. 276, C1361–C1373, 1999. 136. Wang, X.G., Li, L.Q. and Peracchia, C., Chimeric evidence for a role of the connexin cytoplasmic loop in gap junction channel gating, Pflu¨gers Arch. 431, 844–852, 1996.
64
Teppema and Dahan
137. Wang, X.G. and Peracchia, C., Connexin 32/38 chimeras suggest a role for the second half of the inner loop gap junction by low pH, Am. J. Physiol. 271, C1743–C1749, 1996. 138. Ballantyne, D. and Scheid, P., Mammalian brainstem chemosensitive neurons: Linking them to respiration in vitro, J. Physiol. (Lond.) 525, 567–577, 2000. 139. Dean, J.B., Ballantyne, D., Cardone, D.L., Ehrlichman, J.S. and Solomon, I.C., Role of gap junctions in CO2 chemoreception and respiratory control, Am. J. Physiol. Lung Cell Mol. Physiol. 283, L665–L670, 2002. 140. Dean, J.B., Kinkade, E.A. and Putnam, R.W., Intracellular pH and cell-cell coupling in locus coeruleus neurons during mild and severe hypercapnic acidosis, FASEB J. 16, A812, 2002. 141. Saez, J.C., Connor, J.A., Spray, D.C. and Bennett, M.V.L., Hepatocyte gap junctions are permeable to the second messenger innositol, 1,4,5-triphosphate, and to calcium ions, Proc. Natl. Acad. Sci. USA 86, 2708–2712, 1989. 142. Ransom, B. and Sontheimer, H., The neurophysiology of glial cells, J. Clin. Neurophysiol. 9, 224–251, 1992. 143. Deitmer, J.W. and Rose, C.R., pH regulation and proton signalling by glial cells, Progr. Neurobiol. 48, 73–103, 1996. 144. Erlichman, J.S., Li, A. and Nattie, E.E., Ventilatory effects of glial dysfunction in a rat brain stem chemoreceptor region, J. Appl. Physiol. 85, 1599–1604, 1998. 145. Holleran, J., Babbie, M. and Erlichman, J.S., Ventilatory effects of impaired glial function in a brain stem chemoreceptor region in the conscious rat, J. Appl. Physiol. 90, 1539–1547, 2001. 146. Ransom, B., Glial modulation of neuronal excitability mediated by extracellular pH: A hypothesis, Neuroscience 77, 723–743, 1997. 147. Onimaru, H., Arata, A. and Homma, I., Firing properties of respiratory rhythm generating neurons in the absence of synaptic transmission in rat medulla in vitro, Exp. Brain Res. 76, 530–536, 1989. 148. Dean, J.B., Lawing, L.W. and Millhorn, D.E., CO2 decreases membrane conductance and depolarizes neurons in the nucleus tractus solitarii, Exp. Brain Res. 76, 665–661, 1989. 149. Lesage, F. and Lazdunski, M., Molecular and functional properties of twopore-domain potassium channels, Am. J. Physiol. Renal Physiol. 279, F793– F801, 2000. 150. Lesage, F., Pharmacology of neuronal background potassium channels, Neuropharmacology 44, 1–7, 2003. 151. Talley, E.M., Lei, Q., Sirois, J.E. and Bayliss, D.A., TASK-1, a two-pore domain Kþ channel, is modulated by multiple neurotransmitters in motoneurons, Neuron. 25, 399–410, 2000. 152. Washburn, C.P., Sirois, J.E., Talley, E.M., Guyenet, P.G. and Bayliss, D.A., Serotonergic raphe neurons express TASK channel transcripts and a TASKlike pH- and halothane-sensitive Kþ conductance, J. Neurosci. 22, 1256–1265, 2002. 153. Bayliss, D.A., Talley, E.M., Sirois, J.E. and Lei, Q., TASK-1 is a highly modulated pH-sensitive ‘leak’ Kþ channel expressed in brainstem respiratory neurons, Respir. Physiol. 129, 159–174, 2001.
Central Chemoreceptors
65
154. Buckler, K.J., Williams, B.A. and Honore´, E., An oxygen-, acid- and anaesthetic-sensitive TASK-like background potassium channel in rat arterial chemoreceptor cells, J. Physiol. (Lond.) 525, 135–142, 2000. 155. Millar, J.A., Baratt, L., Southan, A.P., Page, K., Fyffe, R.E., Robertson, B. and Matthie, A., A functional role for the two-pore-domain potassium channel TASK-1 in cerebellar granule neurons, Proc. Natl. Acad. Sci. USA 97, 3614–3648, 2000. 156. Patel, A.J., Honore, E., Lesage, F., Fink, M., Romey, G. and Lazdunski, M., Inhalational anesthetics activate two-pore-domain background Kþ channels, Nat. Neurosci. 2, 422–426, 1999. 157. Patel, A.J. and Honore, E., Anesthetic-sensitive 2P domain Kþ channels, Anesthesiology 95, 1013–1021, 2001. 158. Sirois, J.E., Lei, Q., Talley, E.M., Lynch, C., III and Bayliss, D.A., The TASK-1 two-pore domain Kþ channel is a molecular substrate for neuronal effects of inhalation anesthetics, J. Neurosci. 20, 6347–6354, 2000. 159. Van den Elsen, M., Dahan, A., deGoede, J., Berkenbosch, A. and Van Kleef, J., Influences of subanesthetic isoflurane on ventilatory control in humans, Anesthesiology 83, 478–490, 1995. 160. Van den Elsen, M., Dahan, A., deGoede, J., Berkenbosch, A., Van Kleef, J. and Olievier, I.C.W., Does subanesthetic halothane affect the ventilatory response to acute isocapnic hypoxia in healthy volunteers? Anesthesiology 81, 860–867, 1994. 161. Dahan, A., van den Elsen, M.J.L., Berkenbosch, A., DeGoede, J., Olievier, I.C.W., Van Kleef, J. and Bovill, J.G., Effects of subanesthetic halothane on the ventilatory response to hypercapnia and acute hypoxia in healthy volunteers, Anesthesiology 80, 727–738, 1994. 162. Teppema, L.J., Nieuwenhuijs, D., Sarton, E., Romberg, R., Olievier, C.N., Ward, D.S. and Dahan, A., Antioxidants prevent depression of the acute hypoxic ventilatory response by subanaesthetic halothane in men, J. Physiol. (Lond.) 544, 931–938, 2002. 163. Filosa, J.A., Dean, J.B. and Putnam, R.W., Role of intracellular and extracellular pH in the chemosensitive response of rat locus coeruleus neurones, J. Physiol. (Lond.) 541, 493–509, 2002. 164. Wellner-Kienietz, M.C., Shams, H. and Scheid, P., Contribution of Ca2þactivated Kþ channels to central chemosensitivity in cultivated neurons of fetal rat medulla, J. Neurophysiol. 79, 2885–2894, 1998. 165. Pedersen, K.A., Jorgensen, N.K., Jensen, B.S. and Olesen, S.P., Inhibition of the human intermediate-conductance Ca2þ-activated Kþ channel by intracellular acidification, Pflu¨gers Arch. 440, 153–156, 2000. 166. Schubert, R., Krien, U. and Gagov, H., Protons inhibit the BK(Ca) channel of rat small artery smooth muscle cell, J. Vasc. Res. 38, 30–38, 2001. 167. Filosa, J.A. and Putnam, R.W., Multiple targets of chemosensitive signalling in locus coeruleus neurons: Role of Kþ and Ca2þ channels, Am. J. Physiol. 284, C145–C155, 2003. 168. Jiang, C., Xu, H., Cui, N. and Wu, J., An alternative approach to the identification of respiratory central chemoreceptors in the brainstem, Respir. Physiol. 129, 141–157, 2001.
66
Teppema and Dahan
169. Bernards, J.A., Dejours, P. and Lacasse, M.A., Ventilatory effects in man of successively CO2-free, CO2-enriched and CO2-free gas mixtures with low, normal or high oxygen concentrations, Respir. Physiol. 1, 390–397, 1966. 170. Miller, J.P., Cunningham, D.J.C., Lloyd, B.B. and Young, J.M., The transient respiratory effects in man of sudden changes in alveolar CO2 in hypoxia and hyperoxia, Respir. Physiol. 20, 17–34, 1974. 171. Fitzgerald, R.S. and Parks, D.C., Effect of hypoxia on carotid chemoreceptor response to carbon dioxide in cats, Respir. Physiol. 12, 218–229, 1971. 172. Fitzgerald, R.S. and Deghani, G.A., Neural responses of the cat carotid and aortic bodies to hypercapnia and hypoxia, J. Appl. Physiol. 52, 596–601, 1982. 173. Lahiri, S. and Delaney, R.G., Stimulus interaction in the response of carotid body single afferents, Respir. Physiol. 24, 249–266, 1955. 174. Kiwull, P. and Kiwull-Scho¨ne, H., The significance of carotid chemoreceptor stimulus-impulse transmission for the respiratory control system of the rabbit, in Central Neurone Environment and the Control System of Breathing and Circulation, Schlaefke, M.E., Koepchen, H.P. and See, W.R., eds., Berlin, Springer, pp. 102–108, 1983. 175. Schlaefke, M.E., Elimination of central chemosensitivity by coagulation of a bilateral area on the ventral medullary surface in awake cats, Pflu¨gers Arch. 379, 231–241, 1979. 176. Schlaefke, M.E., See, W.R., Herker-See, A. and Loeschcke, H.H., Respiratory response to hypercapnia and hypoxia after elimination of central chemosensitivity, Pflu¨gers Arch. 381, 241–248, 1979. 177. Nattie, E.E., Mills, J.W., Ou, L.C. and St. John, W.M., Kainic acid on the rostral ventrolateral medulla inhibits phrenic output and CO2 sensitivity, J. Appl. Physiol. 65, 1525–1534, 1988. 178. Nattie, E.E. and Li, A.H., Fluorescence location of RVLM kainate microinjections that alter the control of breathing, J. Appl. Physiol. 68, 1157–1166, 1990. 179. Nattie, E.E., Blanchford, C. and Li, A.H., Retrofacial lesions: Effects on CO2-sensitive phrenic and sympathetic nerve activity, J. Appl. Physiol. 73, 1317–1325, 1992. 180. Schlaefke, M.E. and Loeschcke, H.H., Lokalisation eines und der Regulation von Atmung und Kreislauf beteiligten Gebietes und der ventrale Oberfla¨che der Medulla Oblongata durch Ka¨lteblockade, Pflu¨gers Arch. 297, 201–220, 1967. 181. Millhorn, D.E., Eldridge, F.L. and Waldrop, T.G., Effects of medullary area I(s) cooling on respiratory response to chemoreceptor inputs, Respir. Physiol. 49, 23–29, 1982. 182. Budzinska, K., von Euler, C., Kao, F.F., Pantaleo, T. and Yamamoto, Y., Effects of graded focal cold block in rostral areas of the medulla, Acta. Physiol. Scand. 124, 329–340, 1985. 183. Chonan, T., Hida, W., Okabe, S., Izumiyama, T., Sakurai, M. and Takashima, T., Effects of cooling of the ventral medullary surface on breathing pattern and blood pressure in dogs, Respir. Physiol. 83, 77–86, 1991. 184. Homma, I., Isobe, A., Iwase, M., Kanamaru, M. and Sibuya, M., Two different types of apnea induced by focal cold block of ventral medulla in rabbits, Neurosci. Lett. 22, 41–45, 1988.
Central Chemoreceptors
67
185. Forster, H.V., Ohtake, P.J., Pan, L.G. and Lowry, T.F., Effect on breathing of surface ventrolateral medullary cooling in awake, anesthetized and asleep goats, Respir. Physiol. 110, 187–197, 1997. 186. Forster, H.V., Pan, L.G., Lowry, T., Feroah, T., Gersham, W.M., Whaley, A.A., Forster, M.M. and Sprtel, B., Breathing of awake goats during prolonged dysfunction of caudal M ventrolateral medullary neurons, J. Appl. Physiol. 84, 129–140, 1998. 187. Serra, A., Brozoski, D., Hedin, N., Franciosi, R.A. and Forster, H.V., Mortality after carotid body denervation in rats, J. Appl. Physiol. 91, 1298-1306, 2001. 188. Berkenbosch, A., Van Dissel, J., Olievier, C.N., deGoede, J. and Heeringa, J., The contribution of the peripheral chemoreceptors to the ventilatory response to CO2 in anaesthetized cats during hyperoxia, Respir. Physiol. 37, 381–390, 1979. 189. Smith, P.G. and Mills, E., Restoration of reflex ventilatory response after removal of carotid bodies in the cat, Neuroscience 4, 573–580, 1980. 190. Berger, A.J., Krasney, J.A. and Dutton, R.E., Respiratory recovery from CO2 breathing in intact and chemodenervated awake dogs, J. Appl. Physiol. 35, 35–41, 1978. 191. Rodman, J.R, Curran, A.K., Henderson, K.S., Dempsey, J.A. and Smith, C.A., Carotid body denervation in dogs: Eupnea and the ventilatory response to hyperoxic hypercapnia, J. Appl. Physiol. 91, 328–335, 2001. 192. Pan, L.G., Forster, H.V., Martino, P., Strecker, P.J., Beales, J., Serra, A., Lowry, T.F., Forster, M.M. and Forster, A.L., Important role of carotid afferents in control of breathing, J. Appl. Physiol. 85, 1299–1306, 1998. 193. Kiwull, P., Wiemer, W. and Scho¨ne, H., The role of the carotid chemoreceptors in the CO2-hyperpnea under hyperoxia, Pflu¨gers Arch. 336, 171–186, 1972. 194. Forster, H.V., Plasticity in the control of breathing following sensory denervation, J. Appl. Physiol. 94, 784–794, 2003. 195. Katsaros, B., Evidence for the existence of a respiratory drive of unknown origin conducted in the carotid sinus nerves, in Arterial Chemoreceptors, Torrance, R.W., ed., Oxford, UK, Blackwell Scientific, pp. 357–372, 1968. 196. Cohen, M.I., Tonic chemoreceptor input as the background for respiratory rhythm, in Ventral Brainstem Mechanisms and Control of Respiration and Blood Pressure, Trouth, C.O., Millis, R.M. and Kiwull-Scho¨ne, H., eds., New York, Dekker, pp. 797–799, 1995. 197. Hoop, B., Masjedi, M.R., Shih, V.E. and Kazemi, H., Brain glutamate metabolism during hypoxia and peripheral chemodenervation, J. Appl. Physiol. 69, 147–154, 1990. 198. Heeringa, J., Berkenbosch, A., DeGoede, J. and Olievier, C.N., Relative contribution of central and peripheral chemoreceptors to the ventilatory response to CO2 during hyperoxia, Respir. Physiol. 37, 365–379, 1979. 199. Smith, C.A., Saupe, K.W., Henderson, K.S. and Dempsey, J.A., Ventilatory effects of specific carotid body hypocapnia in dogs during wakefulness and sleep, J. Appl. Physiol. 79, 689–699, 1995.
68
Teppema and Dahan
200. Pedersen, M.F., Fatemian, M. and Robbins, P.A., Identification of fast and slow ventilatory responses to carbon dioxide under hypoxic and hyperoxic conditions in humans, J. Physiol. (Lond.) 521, 273–287, 1999. 201. Dahan, A., deGoede, J., Berkenbosch, A. and Olievier, I.C.W., The influence of oxygen on the ventilatory response to CO2 in man, J. Physiol. (Lond.) 428, 485–499, 1990. 202. Dempsey, J. and Forster, H.V., Mediation of ventilatory adaptations, Physiol. Rev. 62, 262–346, 1982. 203. Schuitmaker, J.J., Berkenbosch, A., deGoede, J. and Olievier, C.N., Ventilatory responses to respiratory and metabolic acid-base disturbances in cats, Respir. Physiol. 67, 69–83, 1987. 204. Ahmad, H.R. and Loeschcke, H.H., Fast bicarbonate-chloride exchange between plasma and brain extracellular fluid at maintained PCO2, Pflu¨gers Arch. 295, 300–305, 1982. 205. Teppema, L.J., Barts, P.W.J.A. and Evers, J.A.M., Effects of metabolic arterial pH changes on medullary ECF pH, CSF pH and ventilation in peripherally denervated cats with intact blood brain barrier, Respir. Physiol. 58, 123–136, 1984. 206. Mitchell, R.A., Cerebrospinal fluid pH and the regulation of respiration, in Advances in Respiratory Physiology, Caro, C.G., ed., Baltimore, Williams & Wilkins, pp. 1–47, 1966. 207. Ohtake, P.J., Forster, H.V., Pan, L.G., Lowry, T.F., Korducki, M.J., Aaron, E.A. and Weiss, E.M., Ventilatory responses to cooling the ventrolateral medullary surface of awake and anesthetized goats, J. Appl. Physiol. 78, 247–257, 1995. 208. Nattie, E.E. and Li, A.H., Retrotrapezoid nucleus lesions decrease phrenic activity and CO2 sensitivity in rats, Respir. Physiol. 97, 63–77, 1994. 209. Pearce, R.A., Stornetta, R.L. and Guyenet, P., Retrotrapezoid nucleus in the rat, Neurosci. Lett. 101, 138–142, 1989. 210. Connelly, C.A., Ellenberger, H.H. and Feldman, J.L., Respiratory activity in retrotrapezoid nucleus in cat, Am. J. Physiol. 258, L33–L44, 1990. 211. Ohtake, P.J., Forster, H.V., Pan, L.G., Lowry, T.F., Korducki, M.J. and Whaley, A.A., Effects of cooling the ventrolateral medulla on diaphragm activity during NREM sleep, Respir. Physiol. 104, 127–135, 1996. 212. Forster, H.V., Ohtake, P.J., Pan, L.G., Lowry, T.F., Korducki, M.J., Aaron, E.A. and Forster, A.L., Effects on breathing of ventrolateral medullary cooling in awake goats, J. Appl. Physiol. 78, 258–265, 1995. 213. Spengler, C.M., Gozal, D. and Shea, S.A., Chemoreceptive mechanisms elucidated by studies of congenital central hypoventilation syndrome, Respir. Physiol. 129, 247–255, 2001. 214. Gozal, D., Congenital central hypoventilation syndrome: An update, Pediatr. Pulmonol. 26, 273–282, 1998. 215. Marcus, C.L., Livingston, F.R., Wood, S.E. and Keens, T.G., Hypercapnic and hypoxic ventilatory responses in parents and siblings of children with congenital central hypoventilation syndrome, Am. Rev. Respir. Dis. 144, 136–140, 1991.
Central Chemoreceptors
69
216. Harper, R.M., Gozal, D., Bandler, R., Spriggs, D., Lee, J. and Alger, J., Regional brain activation in humans during respiratory and pressure challenges, Clin. Exp. Pharmacol. Physiol. 25, 483–486, 1998. 217. Spriggs, D., Saeed, M.M., Alger, J.R., Woo, M.A., Woo, M.S., Gozal, D., Keens, T.G. and Harper, R.M., Time course of functional magnetic resonance imaging signal changes in response to hypercapnia in congenital central hypoventilation syndrome, Soc. Neurosci. 25, A111–A112, 1999. 218. Morrell, M.J., Heywood, P., Moosavi, S.H., Guz, A. and Stevens, J., Unilateral focal lesions in the rostrolateral medulla influence chemosensitivity and breathing measure during wakefulness, sleep and exercise, J. Neurol. Neurosurg. Psychiatry 67, 637–645, 1999. 219. Morrell, M.J., Heywood, P., Moosavi, S.H., Stevens, J. and Guz, A., Central chemosensitivity and breathing asleep in unilateral medullary lesion patients: Comparisons to animal data, Respir. Physiol. 129, 269–277, 2001. 220. Grzanna, R. and Fritschy, J.M., Efferent projections of different subpopulations of central noradrenaline neurons, Prog. Brain Res. 88, 89–101, 1991. 221. Haxhiu, M.A., Tolentino-Silva, F., Pete, G. and Mack, S.O., Monoaminergic neurons, chemosensation and arousal, Respir. Physiol. 129, 191–201, 2001. 222. Jacobs, B.L. and Azmitia, E.C., Structure and function of the brain serotonin system, Physiol. Rev. 72, 165–229, 1992. 223. Jacobs, B.L. and Fornall, C.A., An integrative role for serotonin in the central nervous system, in Behavioral State Control: Cellular and Molecular Mechanisms, Lydic, R., Babhdoyan, H.A., eds., Boca Raton, CRC Press, pp. 181–194, 1999. 224. Haxhiu, M.A., Erokwu, B.O., Bhardwaj, V. and Dreshaij, I.A., The role of the medullary raphe nuclei in regulation of cholinergic outflow to the airways, J. Auton. Nerv. System 69, 64–71, 1998. 225. Dempsey, J.A., Smith, C.A., Harms, C.A., Chow, C.M. and Saupe, K.W., Sleep-induced breathing instability, Sleep 19, 236–247, 1996. 226. Malhotra, A., Pillar, G., Fogel, R., Beauregard, J. and White, D., Genioglossal but not palatal muscle activity relates closely to pharyngeal pressure, Am. J. Respir. Crit. Care. Med. 162, 1058–1062, 2000. 227. Nattie, E.E., Central, chemosensitivity, sleep and wakefulness, Respir. Physiol. 129, 257–268, 2001.
3 Suprapontine Control of Breathing
SHAKEEB H. MOOSAVI
DAVID PAYDARFAR
Imperial College of London Faculty of Medicine, NHLI London, UK
University of Massachusetts Medical School Worcester, Massachusetts
STEVEN A. SHEA Harvard Medical School Division of Sleep Medicine Medical Chronobiology Program Brigham and Women’s Hospital Boston, Massachusetts
I.
Introduction
The current chapter focuses on the functional importance and neuroanatomical basis of suprapontine control of breathing in humans, which complements the pharmacological and pathological perspectives covered in other sections of this book. For this purpose, suprapontine influences are divided into three broad categories: (1) volitional control; (2) involuntary influences associated with emotions and psychological disturbances, and (3) tonic drives including excitatory drives associated with wakefulness. Recent reviews in suprapontine respiratory control focus primarily on functional aspects [1–3], or primarily on the underlying neural substrate [4–6]. This chapter updates these reviews with particular regard to recent studies in humans. We focus more on the second category (involuntary influences), as this has never been reviewed extensively. We have also highlighted two key issues that continue to guide research in the field: (1) interaction between suprapontine- and brainstem-based mechanisms that influence respiratory pump activity, and (2) capacity for learning and adaptation in respiratory control. 71
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Definitions and Terminology
Suprapontine control of breathing refers to any influence on the amount or pattern of breathing resulting from neural drives to the respiratory pump muscles that originate in areas of the brain rostral to the pons. Suprapontine control of respiratory muscle activity does not primarily serve metabolic needs and is anatomically distinct from the automatic respiratory controller in the bulbopontine region of the brainstem (selfsustaining respiratory pattern generating neural circuits that automatically adapt to changes in arterial blood gases, thereby subserving metabolic homeostasis). It is convenient to use the functional term behavioral control as a synonym for the anatomical term suprapontine control (and this is distinct from the functional term automatic control which is used as a synonym for the anatomically defined brainstem respiratory controller). In reality, the functional/anatomical distinction is not absolutely precise [7] as some forms of rudimentary behaviors can be elicited by both suprapontine and brainstem sites [8]. These include protective maneuvers and adjustments to breathing necessary to coordinate gustatory brainstem reflexes, which can also be initiated volitionally (e.g., coughing in the absence of irritation) or modulated through reticular mechanisms of arousal [9]. Other synonyms used for the term ‘suprapontine’ include: forebrain, higher-center, non-metabolic, non-automatic and volitional. However, in this chapter we only use the term ‘volitional control’ in regard to ‘willful’ control. III.
Volitional Control
Adult humans can willfully breathe considerably more than is necessary to meet resting metabolic demands. When breathing as fast and deep as possible, ventilation can increase to as much as 40 times resting ventilation. Full recruitment of all inspiratory motor units is possible through willful control [10]. Breathing also can be completely interrupted by willful suppression. When breath-holding from Functional Residual Capacity (FRC), PaO2 will drop by as much as 50 torr and PaCO2 will rise by about 10 torr within 35 sec of breathing cessation [11]. Increased chemoreceptor afferent activity, reflex brainstem respiratory drive and a sensation of an urge to breathe will irresistibly override the voluntary suppression of respiratory muscle activity beyond this time. Starting the breath-hold at a higher lung volume [12], breathing elevated FiO2, hyperventilating beforehand, or practice will all increase breathhold durations by 2–3 min at best. The behavioral advantage conferred by the ability to completely override metabolic demands (even for only a short time) is obvious. Swallowing can occur, breathing muscles can
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be used to alter abdominal pressure necessary for defecation and parturition, noxious fumes can be avoided while moving to a safer environment, sprinters can achieve faster running speeds, and swimming underwater is possible. Why we are able to breathe willfully so much more than we need is less obvious. The maximum voluntary ventilation achievable is approximately 200 l/min, which provides a generous degree of metabolic freedom to accomplish most behaviors requiring willful control. Willful increases in breathing usually have little to do with blood gas and pH homeostasis and may instead incur additional metabolic cost [13]. However, in healthy subjects, ventilation would have to be as high as 140 l/min before any further increase results in a metabolic cost that exceeds the energy potential of the added oxygen transfer [14]. Maximum Voluntary Ventilation (MVV) can only be sustained for a short time (530 sec) before respiratory muscle fatigue sets in [15]. If isocapnia is maintained, as much as 70% of the MVV can be sustained for 4 min [16] and about 50%–60% of the initial MVV can be sustained as long as motivation continues [16,17]. As well as being able to increase and suppress breathing at will, we can also produce precisely guided breathing motion that simultaneously fulfills behavioral as well as metabolic needs. The ability to sing, speak or play musical instruments demonstrates a high degree of precision with which voluntary changes in the amount and pattern of breathing can be manifested. The respiratory pump is essentially a skeletal muscle system, as such willful contractions can be made with response times within 100 msec [18,19]. Volitional tracking of ventilation is as accurate as a control motor task (hand control in operating a joystick) when the rate of tracking is below 20 perturbations per minute [20,21]. In healthy subjects, this degree of control permits regulation of airway pressures to within 2 cm H2O, about 1% of that generated by maximal voluntary effort [22]. Willful control of breathing can become automated as with other rhythmic motor acts (e.g., walking) [23]. Control of breathing for the purpose of speech production exemplifies this form of subconscious suprapontine control. Both metabolic and behavioral needs are usually met without the need to consciously attend to breathing. In a healthy individual, the majority of breaths during wakefulness occur without awareness. However, awareness of breathing increases in the presence of an obstruction to breathing (e.g., nasal congestion); when metabolic demands increase (e.g., heavy exercise); if gas exchange is compromised (e.g., cardiopulmonary disease), or if attempts are made to measure breathing in the laboratory environment. This awareness can itself influence breathing, presenting a substantial confounding factor in experimental procedures. Minimizing the influence requires careful control of measurement procedures, for example avoiding the use of breathing apparatus that can direct attention towards breathing [3].
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There are conflicting messages in the literature about the role of voluntary drives to breathe in patients with severe lung disease. Early studies led to the proposal that patients with increased inspiratory load suffer respiratory failure when the energy consumption of respiratory muscles reaches a critical value, triggering muscle fatigue [24]. Other evidence suggested respiratory failure was due to blunting of voluntary or automatic drives to breathe, perhaps to compensate for or delay the onset of respiratory muscle fatigue [25]. More recent studies paint a somewhat different picture. The ability to track perturbations in ventilation, as compared with the ability to track perturbations in a hand control task, is not compromised in patients with COPD [26]. Healthy subjects fail to breathe to target mouth pressures against inspiratory resistive loads because of dyspnea associated with carbon dioxide retention [10,27,28]. Task failure was not associated with reduction in twitch amplitude of the diaphragm in response to bilateral phrenic nerve stimulation [10,27], whereas earlier studies had assumed that voluntary cessation of the breathing task (task failure) was indicative of respiratory muscle fatigue. Changes in diaphragmatic activity during transient maximal inspiratory pressure maneuvers indicate that the ability to drive breathing volitionally is actually increased in COPD patients with chronic hypercapnia [29,30]. A shift towards volitional drive during loaded breathing may confer some advantage, since volitional control can fully activate all motoneurons to inspiratory muscles whereas automatic control may not be capable of optimal recruitment during maximal reflex stimulation of the bulbopontine controller by chemoreceptor afferents [10]. Voluntary alterations in airflow profile (e.g., slowed expiration) or in end-expiratory lung volume may also increase gas exchange capability. Life without the ability to exercise volitional control over the respiratory apparatus is dramatically illustrated by a rare clinical condition referred to as the locked-in syndrome. A discrete lesion of the corticospinal pathway in the ventral pons [31] characterizes the condition. The usual level of irregularity of the respiratory rhythm during wakefulness is lost (the breathing pattern when awake becomes more like that normally seen in deep non-rapid eye movement sleep) and a reduction in PCO2 of as little as 1 mmHg from normocapnia results in apnea [32], demonstrating regulation of breathing solely for metabolic demands. The decision to take a breath or expire volitionally has its neural origin in the motor cortex. The earliest recognition of this was during neurosurgery in patients under local anesthesia; stimulation of a focal site in the primary motor cortex close to the vertex resulted in contraction of the diaphragm producing a hiccough [33]. The introduction of transcranial electrical stimulation [34] made it possible to demonstrate pathways from the cortex to the diaphragm in normal man [35]. Later, the use of Transcranial Magnetic Stimulation (TMS) [36] confirmed
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that the optimal site of stimulation was a few centimeters anterior and lateral to the vertex [37] at a site consistent with the observations of Foerster [33]. Studies employing Positron Emission Tomography (PET) to detect changes in regional Cerebral Blood Flow (rCBF) (as an index of neuronal activity), have identified specific foci of increased rCBF in the primary motor cortex, the premotor area, the supplementary motor area and the cerebellum during volitional breathing [38–40]. More recent studies employing functional Magnetic Resonance Imaging (fMRI) have verified these activation sites and identified additional cortical and sub-cortical regions that may be involved in mediating a conscious decision to breathe (inferolateral sensorimotor cortex, striatum, thalamus, globus pallidum, lentiform and caudate nuclei, and medulla) [41–43]. Consistency in cerebral activation sites between studies from independent labs and between studies using different imaging techniques strengthens the findings. However, there is uncertainty about the precise role played by individual foci of activations in the programming and execution of voluntary breathing. What is becoming clear is that a distributed neural network is necessary, the extent of which will depend on the degree of learning, planning, attention and concentration involved in the task. The pattern of cortical/subcortical neural activations associated with voluntary breathing resembles that which would be seen with volitional movements in general. The sensitivity and temporal resolution of fMRI now appears to be capable of distinguishing between clusters of activations within the superior motor cortex that are associated with diaphragmatic contraction (anterior) and clusters associated with thoracic muscle contractions (posterior) [43]. Clinical evidence suggests that volitional control involves corticospinal and/or corticobulbar pathways [44]. Studies using TMS to investigate volitional activation of breathing muscles in patients with cerebral ischemia have shown that the cortico-respiratory projections are located in the pyramidal tracts [45] (see also Section VI). IV.
Involuntary Emotional Influences
The study of breathing in relation to emotions spans more than a century and has traditionally been the preserve of psychologists and psychoanalysts (for a thorough and critical review of the extensive body of work see [46]). Our aim in this section is to highlight key observations underlying current notions regarding the functional role of emotion-related breathing modifications. For convenience, we have separated breathing behaviors associated with centrally generated emotions (i.e., psychological factors including depression and anxiety, and higher mental functions such as
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thoughts and memory), from those resulting from the emotional consequences on breathing of ascending sensory afferent information (respiratory and non-respiratory). We recognize that this distinction is somewhat arbitrary since perception of sensory afferent information is likely to influence breathing through psychological disturbance, and conversely, psychological disturbance may modulate perceptual sensitivity [47]. Chemoreceptor and mechanoreceptor afferent activity reaches the suprapontine brain with information about respiration, either directly or via the automatic respiratory controller in the brainstem. These signals provide information about motion, position, local irritation of the respiratory apparatus, blood gas tensions, and sensations of respiratory discomfort (dyspnea) [48]. In addition, a constant barrage of nonrespiratory information is available including sound, sight, smell, taste and pain. Automatic reflex responses can be triggered by direct projection of respiratory (and non-respiratory) sensory afferent activity to structures within the brainstem. For instance, local irritation of the upper airways can trigger a cough to clear the airways. A non-respiratory example is a sudden loud noise, which may elicit a sharp intake of breath. Such reflex behaviors fall outside our definition of suprapontine control; reviews of this form of behavioral control can be found elsewhere (e.g., [8]). We limit our discussion in this section to volitional or emotional influences on respiratory muscle activity that are consequent to psychological state or perceptual processing of sensory information in higher centers. Sensory afferent information may also be involved in maintaining the wakefulness drive to breathe (Section V). A. Psychological Influences
Many emotions are expressed by changes in breathing; for example, laughter normally expresses happiness and sobbing expresses sadness. Less obvious changes can occur; for example, deception is associated with significantly shorter breaths preceding a dishonest response when compared with those preceding an honest response [49]. Various methods have been used to evoke specific emotions in healthy subjects including: self-induction by recall of previously experienced emotions [50], suggestion or hypnosis [51,52], cued presentations of affect-laden words or pictures [53,54] and introduction of artificially created threatening situations [55,56]. In addition, psychiatric assessments of subjective responses to interview have been used to correlate breathing patterns with differing thoughts or moods in the clinical setting [57,58]. Reported breathing modifications include changes in amount or pattern of breathing (see [46] for the various patterns of change reported), changes in the ratio of inspiratory to expiratory durations [50], changes in within-breath parameters [59], changes
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in coordination between ribcage and abdominal motion [60], and irregularities of breathing [61]. Are specific breathing patterns associated with specific emotions? This is a central question raised in the earliest studies, and remains a key question. Two early studies generated interest in a specificity model of emotion-breathing association. Changes in breathing pattern tended to be characteristic of the specific self-induced emotion when subjects were asked to revive thoughts of pleasure, pain, anger, disgust, wonder, fear, laughter or hatred [50], or when subjects were asked to revive joy-laughter, sadnesscrying, anger, fear, erotic arousal and tenderness [62]. While a specificity model cannot be discounted [63], other models of association that relate breathing behaviors to groups of emotions based on different affective dimensions (e.g., pleasant-unpleasant, calm-excited, active-passive) or different response requirements (e.g., fight-flight or passive-active coping) may be more relevant but hard to distinguish [46]. In a study that fits with a dimension model, healthy female subjects asked to generate internal thoughts in response to cued presentations of words, responded with increased respiratory frequency when the affective content of the words was stressful, and responded with decreased respiratory frequency when the affective content was relaxing (relative to neutral words) [53]. Other observations fit better with a response requirement model. In one such study, psychotherapeutic interviews of asthmatic subjects were divided into segments that were judged to be indicative of different emotions based on independent psychiatric assessments. The emotions identified were subsequently divided into four different clusters: neutral, distress, giving-up and arousal. Different patterns of breathing modifications emerged for different emotion clusters as well as with personality differences among individuals [57]. Specific breathing behaviors may also distinguish different psychological derangements. For instance, schizophrenic patients breathe more rapidly and shallowly than either control subjects or melancholic patients [65]. More recently, there is a great deal of interest in the nature of breathing associated with anxiety disorders. If patients with hyperventilation syndrome are asked to recall stressful situations specific to their experience, pronounced hyperventilation can be elicited—this think test has been shown to be more effective in identifying hyperventilators than forced hyperventilation provocation tests alone [65]. Hyperventilation is also a feature of a panic attack that characterizes panic disorder. Breathing is more irregular with greater numbers of sighs and an inappropriate tonic hypocapnia in patients with panic disorder when compared with controls [66–68]. Breathing abnormalities in panic disorder patients are now believed to be due, not to an inherently abnormal respiratory controller, but rather to a hypersensitive fear network comprising the amygdala, its projections to the brainstem, the hippocampus and the medial prefrontal
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cortex [69]. It will not be easy to determine the exact pathways involved in panic disorder in the claustrophobic environment of an MRI or PET scanner. It is also notable that the areas activated by anxiety are close or overlapping with areas activated by the perception of a number of different sensations (pain, nausea, air hunger); anxiety may be an element of unpleasant perceptions that is experimentally inextricable. Whether emotional influences on breathing are characteristic of specific emotions, or of clusters of emotions, is only half the story. In terms of functional significance, we must also consider what role such associations might serve. The functional role of psychological influences on breathing is not known. The changes in breathing associated with psychological factors usually occur without awareness and are unrelated to metabolic requirements. Expression of emotions can greatly reduce the ability of the respiratory apparatus to participate in its metabolic role. For instance, fits of laughter produce substantial reductions in lung volume and increase in dynamic airway compression [70]. However, breathing adjustments are not always counterproductive. It is possible that the effects are not simply an epiphenomenon of psychological state but serve some physiological, behavioral or adaptive purpose. Sighs and yawns counter atelectasis of the lungs; such recruitment maneuvers may be especially beneficial in the presence of pulmonary disease [71]. In certain stressful situations, slowing breathing can alleviate anxiety [72]. Subjects consciously performing physical relaxation techniques produce more coordinated ribcage and abdomen motion [60]. Attention and motivation are likely to be necessary for behavioral adaptation of volitional or reflex breathing responses. For instance, attention or motivation can substantially prolong breathholds [73]—being able to breath-hold for longer is a distinct advantage in some circumstances. There is much speculation about the broader impact of emotional influences on breathing, which remains largely unexplored and unsupported by experimental evidence. One possibility is that the changes in breathing brought about by psychological factors are inherently involved in the establishment and self-regulation of emotions. This idea stems from an early theory that physical changes play an important role in formation and experience of emotions [74]. Accordingly, breathing, among other physiological activities, may play a vital role in forming negative emotions that ensure avoidance of harmful situations, thereby contributing to evolutionary survival. Breathing adjustments by psychological factors may also represent a subconscious primeval form of communication; yawns and sighs may communicate boredom and melancholy, but may also indicate contentment [75], and are highly contagious [76]. Interestingly, yawn contagion appears to be found exclusively in humans [77]. What are the cerebral correlates of emotion perception in humans? Within the last decade, a vast literature has built up on this subject
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based on functional neuroimaging, electrophysiological and lesion studies (see [78] for a critical review). In general, there appears to be two neural networks. The emotional impact of a stimulus and its behavioral consequences are processed within a ventral neural network (comprising the amygdala, insula, ventral striatum, and ventral regions of the anterior cingulate and prefrontal cortex). Regulation of the behavioral responses is thought to be the predominant function of the dorsal neural network (comprising dorsal regions of the anterior cingulate gyrus and the prefrontal cortex). What do we know about the descending pathways with respect to emotion-related breathing behavior? Emotion-related breathing behavior can be expressed through involuntary control with no awareness of breathing changes. There is some evidence that this form of influence on the respiratory pump muscles can occur through descending suprapontine pathways that are separate from volitional suprapontine control pathways. For example, there are changes in breathing associated with a display of emotion in patients who have lost volitional control of breathing due to a lesion of the corticospinal tract [31]. Several involuntary motor patterns involving respiratory muscles have been observed in these patients including whining, moaning, sighing and yawning [79], but there is still much to be learned from these unfortunate patients. For instance, it is not yet known if they can learn to use the emotional pathway to enable voluntary activation of the respiratory muscles despite obstruction of corticospinal/corticobulbar pathways. B. Perception of Respiratory Discomfort
Respiratory sensations are always available to the forebrain to direct motor activity; these sensations and the consequent conscious or subconscious suprapontine influences upon breathing may be important in shaping much of our breathing behavior while we are awake. A broader discussion of the influence of respiratory sensations on breathing can be found in earlier volumes in this series [80,81]. Here we have focused specifically on perceptions of respiratory discomfort since much progress has been made in recent years with regard to identifying separable qualities of respiratory discomfort and their most likely afferent sources. Clearly, the framework of perceptual mechanisms for unpleasant breathing sensations has grown in complexity. Physicians refer to sensations of respiratory discomfort as dyspnea (though some use the term in its literal sense, meaning difficult and labored breathing). Patients commonly report dyspnea as shortness of breath or breathlessness. At least three separate qualities of respiratory discomfort can be perceived by patients and healthy subjects: air hunger, effort or work and tightness. This classification is based on studies that analyzed
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the quality of unpleasant sensations evoked by different respiratory stimuli in normal subjects [82,83] or by different clinical conditions in patients [84,85]. Air hunger is readily recognizable as the unpleasant, uncomfortable, imperative urge to breathe at the culmination of a long breath-hold. The sensation of breathing effort or work arises when ventilation requires excessive respiratory muscle activity, e.g., due to fatigue or increased impedance to inspiration [86–89]. The sense of chest tightness is usually associated with episodes of bronchoconstriction [84,90,91]. The various forms of respiratory discomfort coexist in the clinical arena. The relative contribution of each to the overall perception will depend on the underlying pathology. Laboratory evidence shows that these distinguishable sensations arise from different afferent sources [86,87,90,91]. Intense dyspnea can elicit emotional disturbance leading to behavioral adaptations. Dyspnea is a powerful symptom of disease that motivates patients to seek medical attention and to adopt other non-specific changes in lifestyle. Modification of breathing behavior can result in a number of ways. Psychological factors can subconsciously influence breathing behavior directly. As discussed in the previous section, the purpose of such influences is uncertain in some situations, changes may alleviate the perception or the underlying breathing difficulty while in others they may be counterproductive. Dyspnea perception may also directly influence volitional drives to breathe or influence drives to other muscle systems that indirectly influence breathing behavior. These forms of modulation may serve to minimize unpleasant breathing sensations [92]. The impact of dyspnea on breathing behavior will depend on the degree to which the perception is affect-laden. Since air hunger is a particularly unpleasant form of dyspnea, one might expect that this form of dyspnea would more readily influence breathing. Normal subjects and patients with COPD are able to distinguish between the intensity of dyspnea and the distress or anxiety associated with it [93,94]. However, it is yet to be demonstrated that the relationship between unpleasantness and intensity differs among the various qualities of dyspnea, as has been shown for different qualities of pain perception [95]. It is very likely that dyspnea perception will likewise be found to be multi-dimensional in nature [96]. What evidence is there that perception of respiratory discomfort directly influences volitional drives to breathe? It makes intuitive sense that conscious awareness of unpleasant breathing sensations leads individuals to access volitional control pathways in an attempt to minimize the sensation. Based on a growing body of evidence, air hunger depends on excitatory ascending projections of efferent respiratory drives from the brainstem (that report prevailing ventilatory demand) modulated by inhibitory afferent information from pulmonary mechanoreceptors (that report the present level of ventilation). Thus, substituting part of the reflex ventilatory demand from the brainstem with volitional drive may serve to alleviate the urge
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to breathe. It has been proposed that the ventilatory response to hypercapnia may involve a cortical facilitatory component aimed at reducing the urge to breathe that accompanies hypercapnia [97]. This was later challenged by the observation that the minimum ventilation required to alleviate the air hunger of hypercapnia was less than the spontaneous breathing during hypercapnia [98]. Brain imaging studies aimed at elucidating the cerebral activations associated with CO2-stimulated breathing in awake man provide conflicting evidence with regard to the contention that hypercapnia induces cortical facilitation of breathing. Those based on PET studies have found no evidence of primary motor cortex activation [99] in areas previously shown to activate with volitional breathing [39,40]. Other studies based on fMRI techniques suggest that motor cortical activation does accompany the ventilatory response to hypercapnia [100]. Although in some circumstances it would make sense to voluntarily activate breathing muscles for the purpose of minimizing unpleasant respiratory sensations (e.g., terminating a prolonged breath-hold if it is safe to do so), it would be counterproductive in other situations. For instance, a diver must continue to hold his/her breath, despite tremendous metabolic drive, until the swim back to the surface is complete. The decision to begin the swim back to the surface must be made well in advance of intolerable air hunger. Voluntary activation of breathing muscles would also be counterproductive when the ventilatory apparatus is already compromised by disease. This could generate more unpleasant sensations, thereby setting up a positive feedback situation. Thus, it is possible that the perception of air hunger evolved not to directly influence breathing but to generate more general adaptive behaviors serving to distance the individual from situations that threaten respiratory function. A number of behavioral responses exhibited by patients and healthy subjects that may represent subconsciously learned strategies to avoid or minimize unpleasant breathing sensations are discussed in Section VII. Several patient groups have been identified in whom inappropriate perceptions of respiratory discomfort are apparent. An inappropriately high sensitivity of air hunger is a very common symptom in hyperventilation syndrome [101,102], and it is possible that this sensation is itself the trigger for psychogenic hyperventilation. Hyperventilation syndrome may develop as a vicious circle between breathing response and the negative emotional content of symptoms associated with it [103,104]. Increased dyspnea sensitivity to hypercapnia is also apparent in panic disorder; it is uncertain whether this is integral to an inappropriate fear response [69]. At the other end of the spectrum, patients with blunted perception of dyspnea (so-called poor perceivers) may account for many asthma fatalities [105]. A recent study has also reported blunted perception of dyspnea in stroke patients [106]. Depression in chronic fatigue syndrome can increase the effect of work or effort of breathing associated with resistive loading [47] and may
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impair voluntary activation of the diaphragm in asthmatics, making them more vulnerable to ventilatory failure [107]. What are the cerebral correlates of dyspnea perception? Three PET studies [108–110] and one fMRI study [111] have recently mapped brain activations during dyspnea induced in normal subjects. Principal activation was in the insula (extending to the operculum), cerebellum, and the anterior cingulate (other activations included thalamus and SMA). Activation of the anterior insular cortex was common in all of these studies, raising the possibility that the insula is essential for dyspnea perception. The insular cortex is a para-limbic structure. One of its many roles is thought to be formulating behaviors and learning visceral homeostasis [112]. Electrophysiological and tracer studies in animals indicate that it receives afferents from respiratory chemoreceptors, pulmonary receptors and medullary respiratory neurons [113–115]. The insula is activated in imaging studies of other unpleasant visceral sensations [116,117], including pain [118]. What are the different afferent sources of dyspnea perception? The sensation of air hunger (e.g., during blood gas derangements) most likely arises from perception of respiratory motor drive from the brainstem [86,119], a corollary copy of which is transmitted to the cerebral cortex [120,121]. In addition, a direct projection of chemoreceptor afferents cannot be discounted. Respiratory muscle activity is not involved in generating air hunger, since complete neuromuscular blockade does not diminish this sensation [122,123]. Nonetheless, afferent information from respiratory muscle activity and other mechanoreceptor afferent information reporting the level of pulmonary ventilation are known to relieve dyspnea [124,125]. Evidence points to vagal afferent information from pulmonary receptors, activated by bronchospasm or inflammation of the airways, as the most likely source of tightness [90]. The sense of breathing effort or work is thought to arise from a combination of respiratory muscle afferent activity and from corollary discharge of central neural motor drives to the respiratory muscles [86,89]. In contrast to air hunger, pulmonary afferent information is not thought to have a role in relieving sensations of breathing effort or work. C. Perception of Music
Anticipation of experimental procedures [126,127] and states of anxiety [128] can give rise to increased ventilation and/or irregularity of breathing patterns. Respiratory studies on human subjects therefore often employ music or other auditory stimuli (such as white noise or talking stories) in order to distract subjects from the test stimuli or more generally to relax naive subjects or patients who may be anxious. However, a number of studies have shown that auditory stimulation may itself introduce a source of ‘‘behavioral noise’’ that can affect the amplitude and cycle period of
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respiration. Poole et al. [129] demonstrated systematic effects of different tone bursts on the breathing cycle and suggested that such effects could form the basis of objective hearing tests in children or patients with psychological pathologies. Early studies by Skaggs [130] employed novel forms of auditory stimulation (a car horn and an empty can dropped on the floor) to demonstrate, not surprisingly, sudden reactive inspiratory movements. Other studies [131,132] investigated these effects using transient presentations of white noise and showed that the respiratory cycle during which they were presented, as well as subsequent cycles, were affected. Since the effects decayed with repeat presentations, this supports the view that they represent orienting or defensive reflexes to which the subjects habituate [131]. Nonetheless, continued white noise over many respiratory cycles at a comfortable auditory level systematically increases breathing frequency, perhaps via an arousing influence [133]. Indeed, direct stimulation of the reticular activating system may represent a general arousal mechanism through a change in the contribution of a tonic wakefulness drive on respiratory control (see Section V). The above forms of direct stimulation may bear a teleological significance in eliciting fight/flight reflex responses. The effects of musical or rhythmical auditory stimulation over many respiratory cycles have also been reported [134,135]. The close anatomical proximity of auditory nuclei to respiratory neural networks in the brainstem and the abundance of collaterals and interneurons in the auditory pathway make it likely that some form of direct stimulation of the automatic respiratory controller is possible when subjects attend to musical auditory inputs. Haas et al. proposed that music rhythm acts as an external pacemaker (‘zeitgeber’) that ‘modulates’ respiratory timing [135] by impinging on the respiratory central pattern generator (CPG) and entraining breathing. In support of this hypothesis, these authors were able to demonstrate a reduced coefficient of variation in breath period when subjects listened to music as well as to demonstrate integer ratios between beat period (of a metronome or of various musical segments) and breath period; they also demonstrated a significant respiratory phase coupling in 12 out of 20 subjects (half of whom were musically trained). The study by Harrer [134] is often cited as a particularly dramatic example of behavioral modulation of breathing (e.g., cited in [2]). This study demonstrated that, in a human subject listening to a melodic excerpt of music by Chopin, respiratory volume became markedly irregular when the music was suddenly replaced by the dissonant atonal electronic music of Stockhausen. However, we are not aware of any other experimental verification of this single observation. Music (especially classical music) is often touted as a ‘relaxant’ or ‘mood enhancer.’ Music has been shown to reduce anxiety and respiratory rates in women awaiting surgery [136] and reduces spontaneous breathing efforts in ventilator-dependent patients [137]. Functional neuroimaging
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studies suggest that the perceptual processing of the pleasantness or unpleasantness of music involves the paralimbic and neocortical regions [138]. V.
Tonic Excitatory and Inhibitory Drives
Sections III and IV indicate that during the awake state there are at any instance a great many potential influences on breathing. A recurring notion that would explain the subconsciously increased breathing in many different behavioral situations in awake subjects is that the increased breathing is a by-product of arousal. Fink [139] coined the term the wakefulness drive to breathe. In his study, thirteen subjects underwent passive (mechanically ventilated) and active hyperventilation to reduce PaCO2 and increase PaO2. Hence, any ‘chemical’ drive to breathe was absent at the end of either of these periods. But, it was found that rhythmic breathing continued at the normal frequency in all subjects who remained awake. What was keeping the respiratory rhythm going? Fink observed that one of the subjects stopped breathing when he became drowsy. This momentary response in the drowsy subject is the same as occurs in humans under general anesthesia or sleep wherein apnea is elicited consistently by hypocapnia (e.g., [140– 143]). Fink concluded that cerebral activity associated with wakefulness is a component of the normal respiratory drive. However, it has never been proven that it is cerebral activity per se which enhances breathing, and it could represent activity in many of the brainstem and basal forebrain areas that are associated with arousal. For instance, the loosely defined reticular activating system, which embeds the brainstem respiratory complex, can provide a stimulatory effect upon certain brainstem respiratory-related neurons [144,145]. There are other more specific arousal related neurons throughout the brainstem and basal forebrain [146] and it is possible that these stimulate breathing directly. Absence of this tonic drive to breathe at sleep onset can cause hypoventilation and even central apnea (this appears more likely when sleep transitions are rapid). For a comprehensive review of this area, see Shea [3]. In slow-wave sleep, there is a consistent reduction in ventilation, associated with an increase of several mmHg in the arterial PCO2 [147]. Medullary and pontine inspiratory related neurons show a significant decrease in activity [148]. This phenomenon is essentially independent of influences from peripheral sensory input; extensive peripheral denervation does not abolish these sleep-related decrements in the firing of respiratoryrelated neurons. It has been suggested that this decrease in respiratory activity probably represents loss of tonic excitatory activity associated with wakefulness [149]. Support for this idea in humans comes from a study of breathing during ventilator-induced hypocapnia in normal subjects [141].
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During wakefulness, only brief periods of apnea were obtained. However, during sleep, at comparable levels of hypocapnia, apnea lasted longer than in awake subjects. Furthermore, the duration of apnea was longer in deeper stages of non-REM sleep. Hypoxia could shorten the duration of apnea. These and other studies conclusively demonstrate that in slow-wave sleep, breathing is critically dependent upon arterial PCO2 and PO2. Breathing during REM sleep is much more variable and less influenced by chemical drives, and is thought to be essentially controled by the behavioral control system (reviewed in [3]). Therefore, derangements in the automatic or homeostatic control of respiration are manifested mainly during non-REM sleep and may not be obvious during wakefulness or REM sleep. A dramatic example of this is Congenital Central Hypoventilation Syndrome whereby patients breathe normally during wakefulness and REM sleep but seriously hypoventilate during non-REM sleep (see Chapter 9, this volume, and [150]). Animal studies in which different parts of the brain were electrically stimulated, showed that there were more regions providing an inhibitory influence on the automatic respiratory controller than excitatory influences [151]. Seizures are known to elicit prolonged apnea, especially when their focus is in the limbic areas [152]. Conversely, bilateral damage to the hemispheres can ‘release’ involuntary behavioral control of respiratory muscles that are normally inhibited. For example, pathological involuntary laughter or crying can occur if limbic projections to subcortical structures are impaired [7]. Such ‘disinhibition’ is commonly seen as a psychiatric consequence of traumatic brain injury. Cerebral infarcts are also known to result in increased ventilatory sensitivity to hypercapnia [153,154]. Existence of cortical inhibition of diencephalic structures is strongly supported by lesion experiments in animals. Thus the ventilatory response to hypoxia is unchanged by midcollicular section but dramatically enhanced by decortication [155]. The activity of the automatic controller in the brainstem is therefore influenced not only by tonic excitatory influences associated with wakefulness, but also by tonic inhibitory drives that provide cortical braking of ventilatory reflexes. Neuroanatomically, sources of tonic excitatory influences are located in subcortical structures such as the hypothalamus and reticular formation, whereas tonic inhibitory influences descend from more rostral limbic areas serving to modulate the excitatory influences or directly braking activity of the automatic controller in the brainstem. It has been proposed that tonic excitatory and tonic inhibitory influences on the metabolic controller exist in a state of balance that can be shifted in certain circumstances such as hyperthermia, chronic exposure to high altitude and when different doses of certain general anesthetics are given [4]. It has been suggested that cortical inhibition may be a remnant of a diving reflex [156]. Cerebral damping of metabolic reflexes may also play an important role
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in protecting against instability of respiratory control as in Cheyne Stokes breathing [153]. VI.
Interaction between ‘Behavioral’ and ‘Automatic’ Control
The many reasons for which we breathe can be broadly divided into those exclusively serving ‘metabolic’ needs and those primarily serving ‘behavioral’ needs. This functional hierarchy is partly reflected in an anatomical hierarchy. The existence of a separate automatic controller in the brainstem is supported by evidence from a large number of studies using different methodological approaches [157], including tissue ablation [155]. The behavioral system of control is less well defined and more widely distributed, extending from the brainstem to the highest areas of the brain [3,6,8]. The majority of sensory inputs and the final neural output to pump muscles are common to both brainstem and suprapontine control mechanisms. The respiratory pump apparatus is often described as unique among motor systems since two separate controllers drive it concomitantly. However, the uniqueness of the respiratory motor system may simply be that its automatic central pattern generator resides at a lower neural threshold. This viewpoint is indirectly supported by evidence from animal work [158] that implicates gating mechanisms capable of triggering automatic rhythmic muscle contractions for locomotion without cortical input. The neurological basis and extent to which the behavioral and metabolic controls of breathing interact were realized as key issues several decades ago [7,159]. The signal that underlies automatic/metabolic control of breathing is transmitted to spinal motoneurons via ventrolateral tracts in the spinal cord. There is clinical and experimental evidence that suprapontine drives are transmitted to the same group of motoneurons involved in automatic respiration, but via dorsolateral spinal cord pathways [160,161]. However, the pathways within and rostral to the brainstem are less certain. One possibility is that the voluntary signal originates in the contralateral cortex and bypasses the automatic control system in the brainstem. In this model, the integration of the voluntary and automatic signals occurs exclusively at the level of respiratory motoneurons in the spinal cord. However, it has been demonstrated in conscious animals that a learned voluntary respiratory task (abrupt termination of inspiration and prolongation of expiration) was associated with concomitant termination in brainstem inspiratory neurons and activation of some brainstem expiratory neurons [162]. These findings suggest that the voluntary signal may be integrated at least in part at the level of the automatic system. A recent fMRI study has reported the presence of medullary activity associated with voluntary breathing in humans [43], but could not exclude the possibility
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that this activity was related to sensory afferent input from the lungs or chest wall. There are many situations in which the behavioral and metabolic control systems have conflicting influences on breathing and must therefore compete for control over the respiratory pump. It has been suggested that during much of the awake state, breathing is under the exclusive control of suprapontine mechanisms [3,163]. The observation that the precision with which ventilation can be volitionally tracked is the same irrespective of whether a background of hypocapnia or one of eucapnia is maintained, is consistent with the notion that chemical control plays little or no part during resting breathing [21]. Thus, in the awake state, the relative effectiveness of regulating influences will vary depending on the imperativeness of behavioral needs as much as upon metabolic needs [139]. To what extent can volition dictate breathing before metabolic needs become imperative? Duration of maximum voluntary breathing efforts is less with inspiratory resistive loading [24,164] or elevations in lung volume [165]. If isocapnia is not artificially maintained, syncope could occur due to cerebral ischemia—a natural limitation to willful hyperventilation. The duration of breath holds also depends on lung volumes as well as upon blood gas levels. When metabolic demands are increased, as during exercise, the ability to exert willful control on breathing is further curtailed as exemplified by significant reduction in breath hold durations [166] and by a reduction in speech intelligibility [167]. Motivation, which in turn will depend on the level of attention or arousal, emotional state and other psychological factors, plays an important role in determining the degree to which the metabolic controller can be challenged. Thus, the duration of a breath hold will increase substantially in highly motivated situations such as competition. Elite synchronized swimmers can hold their breath for significantly longer durations than control subjects despite the fact that their hypercapnic ventilatory sensitivity is not different [168]. Presumably they can tolerate higher CO2 loads. Dejours [169] pointed out that during 100 m sprints, elite athletes might not breathe at all despite a tremendous metabolic burden, implying that all proposed automatic drives to breathe can be overridden by the cortex at least for a short time. Speech provides a good example of the competition between the behavioral and automatic respiratory control systems. The flow requirements for normal speech exceed resting ventilation, and so subjects hyperventilate when they speak at rest (e.g., [159,170]). However, as the automatic drive is increased, say by exercise, the subject will have some control of whether to maintain their speech quality at the expense of reduced breathing, or to increase the airflow during speech (thereby altering speech quality) to maintain their gas exchange. The first approach appears to predominate—for example, there were 73% reductions in the ventilatory responses to hypercapnia during speech [159,170], and a 55% reduction
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in the ventilatory response to exercise during speech [171]. A more recent study reports similar findings [172]. Nevertheless, at the extremes of automatic ventilatory drive, subjects may not be able to speak because of the high flows required for gas exchange. For example, during very heavy exercise only gasped phrases are possible [167,171]. It should be pointed out that in these studies, the air used for speech has already been used for gas exchange, so the behavioral and automatic respiratory drives are not completely mutually exclusive. However, in ventilator-dependent tracheostomized patients, there is complete antagonism between the airflow requirements for speech and gas exchange; in this situation, air is stolen from alveolar ventilation during the ventilator’s inflation phase to produce sound [173]; thus there is hypoventilation and relative hypercapnia during speech. During induced hypercapnia (15 mmHg increase in PETCO2 ) that caused shortness of breath, all subjects could still speak adequately. Two of five subjects adapted by reducing the air used for speech during inflation. In contrast, one subject reacted, as normal subjects do during hypercapnic speech, by increasing the airflow per syllable (a maladaptive strategy in ventilated subjects, because this causes relatively greater hypoventilation). These adaptive or maladaptive changes were modest despite the strong hypercapnic stimulus. No subject adapted fully, by consistently not speaking on the ventilator’s inflation phase. It was concluded that the behavioral drive to speak was modified but never fully suppressed by increased automatic respiratory drive. VII.
Learned Respiratory Behaviors
Behavioral strategies to avoid respiratory discomfort include stopping exercise, altering eating habits and speech patterns (and possibly language), some of which involve modulation of muscle activity not normally part of the breathing act. Compared with healthy subjects, patients with sarcoidosis, asthma, or emphysema adopt different breathing patterns during conversational speech, thereby avoiding shortness of breath [174]. Nasal congestion is associated with uncomfortable perceptions of resistance to airflow; even in healthy subjects, this alerts the individual to increase respiratory muscle force or to switch to mouth breathing. Patients with severe cardiopulmonary disease often adopt pursed-lips breathing, brace their arms, or elevate functional residual capacity with tonic inspiratory muscle contraction (e.g., [175,176]). Pursed-lips breathing improves some mechanical indices of lung function by promoting slower and deeper breathing [177–179]. Alleviation of dyspnea and improvements in arterial blood gases have been observed in COPD [179], but pursed-lips breathing may not relieve certain forms of dyspnea [177]. Arm bracing allows normal subjects to increase by a small amount the maximal ventilation they can
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sustain [180]; this may explain why athletes brace their arms on their knees at the end of a sprint to repay the oxygen debt. A more recent study has shown that the increased capacity to sustain hyperpnea cannot be explained by improved inspiratory muscle function that is in fact impaired by arm bracing [181]. Learned behavioral respiratory adaptations may be based upon the improvement in performance of a task. The precisely guided voluntary respiratory movements involved in singing or playing wind instruments are an example of learned respiratory adaptation motivated by a desire to improve performance outcome. Feedback of internal respiratory sensations (proprioceptive cues from the lungs and chest wall) as well as external sources of feedback (e.g., auditory) help to shape the complex pattern of efferent motor drives needed. Repetition and practice will reduce the need for mental concentration and attention to feedback, indicating a reduced dependence on cortical motor planning and greater involvement of subcortical structures such as basal ganglia and cerebellum. Experimental verification exists of the idea that a voluntarily altered breathing pattern can be improved with training, especially when there is feedback (in this case electromyographic) [182]. Breathing retraining techniques based on these principles have had some success in rehabilitation of patients with lung disease (e.g., [183,184]) and hyperventilation syndrome [185,186]. The longterm clinical effectiveness of these breathing retraining techniques is as yet inconclusive. It has yet to be determined how long one can retain a voluntarily modified breathing behavior, and whether the learned pattern ever becomes completely automatic. In this respect, sleep may be a useful tool to distinguish automatic from subconsciously controlled breathing patterns. Exercise hyperpnea in humans may be in part a learned response that has been forged by adaptive adjustments [187,188]. This scheme of control would be independent of any explicit exercise input and thereby differs from the idea that the descending motor command to exercising muscles simply irradiates to respiratory neurons in the brainstem [127]. Feedforward control of this type presumably depends on a memory of the prior experience of exercise and might therefore require plasticity in respiratoryrelated neurons within suprapontine neural networks. The possibility that such plasticity can occur at the level of the brainstem cannot be discounted. The first experimental evidence for this hypothesis came from a study in awake goats [189] in which the goats hyperventilated during exercise after they had been subjected to training trials in which the same exercise was performed repeatedly but with added respiratory deadspace in the breathing circuit. Several recent reports have generated conflicting claims about whether such long-term modulation of exercise hyperpnea is possible in humans. One study found greater ventilatory responses at exercise onset following several trials of arm exercise with added respiratory deadspace [190].
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However, a clear demonstration of long-term modulation of the steady-state phase of exercise in humans is lacking [191,192]. It is possible that greater numbers of conditioning trials or more elaborate associative conditioning paradigms may be required to alter a response that has been forged over many years. There have been many attempts to demonstrate learned respiratory behaviors through classical conditioning of respiratory responses to changes in blood gases (see [193] for review of the early work). In a well-controlled study in humans, it has been demonstrated that following eight pairings of a sound and a brief hypoxic challenge, the sound alone produced an increase in breath duration [194]. Cats have been conditioned to hold their breath when presented with a tone by initially presenting the tone in combination with an inhalation of ammonia [162]. The marked changes in breathing which can occur during the anticipation of performing exercise are thought to be due to a learned feedforward response to avoid blood-gas disturbances [126]. Such adaptability of the ventilatory responses to, for example, exercise and hypoxia may be important in maintaining normal blood gases in the face of changes in physiological, mechanical or environmental conditions. VIII.
Summary
Without the various forms of volitional control of breathing we would not be able to sing, speak, whistle, play wind instruments, sniff, swim or perform yoga—activities that define humans both as individuals as well as social beings. Emotional influences on breathing may occur as a result of internally generated psychological disturbances that are pathological (depression, anxiety, panic disorder) or non-pathological (mental stress, thoughts and memory). Emotional influences may also occur through perceptual processing of respiratory or non-respiratory sensory afferent information. Such influences may modulate respiratory as well as nonrespiratory muscle systems. The functional significance of such influences is less certain; possible roles include rudimentary forms of social communication; protection of the respiratory apparatus from threatening situations that cannot be dealt with by simply changing breathing; improving mechanical efficiency of the respiratory apparatus, and in the development of emotions themselves. The operation of the automatic controller in the brainstem is modulated by tonic excitatory influences (from the subcortical structures) as well as tonic inhibitory influences (from more rostral limbic areas). Many of the emotional influences may be by-products of a shift in the balance of these opposing influences. This balance may also serve to stabilize respiratory control. How and to what extent this suprapontine system of control interacts with the automatic system of control and adapts, remain
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key issues in this field. Direct study of these issues in human subjects has benefitted in recent years from rapid advances in functional neuroimaging techniques and non-invasive transcranial stimulation techniques, guided by pathological case reports. Acknowledgments SHM is a Parker B. Francis Fellow in Pulmonary Research. DP is supported by grants from the National Institutes of Health (NIH R01 HL49848, HL071884) and an Established Investigator Award from the American Heart Association (AHA). SAS received support from the National Institutes of Health (NIH R01 HL64815 and NIH K24076446). The contents of this chapter are solely the responsibility of the authors and do not necessarily represent the official views of the NIH or the AHA.
References 1. 2.
3. 4.
5.
6. 7.
8.
9. 10.
Guz, A., Brain, breathing and breathlessness, Respir. Physiol. 109, 197–204, 1997. von Euler, C., Introduction: Forebrain control of breathing behavior, in Respiratory Psychophysiology, Vol. 50, von Euler, C. and Katz-Salamon, M., eds., Basingstoke, Macmillan Press, pp. 1–14, 1988. Shea, S.A., Behavioral and arousal-related influences on breathing in humans, Exp. Physiol. 81, 1–26, 1996. Hugelin, A., Forebrain and midbrain influence on respiration, in Handbook of Physiology, Section 3, The Respiratory System, Vol. II, Cherniack, N.S. and Widdicombe, J.G., eds., Baltimore MD, American Physiological Society, pp. 69–91, 1986. Davenport, P.W. and Reep, R.L., Cerebral cortex and respiration, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, Inc., pp. 365–388, 1995. Horn, E.M. and Waldrop, T.G., Suprapontine control of respiration, Respir. Physiol. 114, 201–211, 1998. Plum, F., Neurological integration of behavioral and metabolic control of breathing, in Breathing: Hering-Breuer Centenary Symposium, Porter, R., ed., London, Churchill, pp. 159–181, 1970. Dick, T.E., Orem, J.M. and Shea, S.A., Behavioral control of breathing, in The Lung, Scientific Foundations, Crystal, R.G. and West, J.B., eds., Philadelphia, Lippincott-Raven, 1996, pp. 1821–1837. Cohen, M.I. and Hugelin, A., Suprapontine reticular control of intrinsic respiratory mechanisms, Arch. Ital. Biol. 103, 317–334, 1965. McKenzie, D.K., Allen, G.M., Butler, J.E. and Gandevia, S.C., Task failure with lack of diaphragm fatigue during inspiratory resistive loading in human subjects, J. Appl. Physiol. 82, 2011–2019, 1997.
92
Moosavi et al.
11.
Sasse, S.A., Berry, R.B., Nguyen, T.K., Light, R.W. and Mahutte, C.K., Arterial blood gas changes during breath-holding from functional residual capacity, Chest 110, 958–964, 1996. Fowler, W.S., Breaking point of breath-holding, J. Appl. Physiol. 6, 539–545, 1954. Cotes, J.E., The ventilatory cost of activity, Br. J. Ind. Med. 32, 220–223, 1975. Otis, A.B., The work of breathing, Physiol. Rev. 34, 449–458, 1954. Tenney, S.M. and Reese, R.E., The ability to sustain great breathing efforts, Respir. Physiol. 5, 187–201, 1968. Freedman, S., Sustained maximum voluntary ventilation, Respir. Physiol. 8, 230–244, 1970. Martin, B., Heintzelman, M. and Chen, H.I., Exercise performance after ventilatory work, J. Appl. Physiol. 52, 1581–1585, 1982. Plassman, B.L. and Lansing, R.W., Perceptual cues used to reproduce an inspired lung volume, J. Appl. Physiol. 69, 1123–1130, 1990. Lansing, R.W. and Meyerink, L., Load compensating responses of human abdominal muscles, J. Physiol. 320, 253–268, 1981. Heywood, P., Murphy, K., Stong, B., Belyavin, A., Farmer, E. and Guz, A., Is corticospinal control of ventilation as precise as control of hand movement? J. Physiol. 452, 30P, 1992. Corfield, D.R., Roberts, C.A., Guz, A., Murphy, K. and Adams, L., Modulation of the corticospinal control of ventilation by changes in reflex respiratory drive, J. Appl. Physiol. 87, 1923–1930, 1999. Draper, M., Ladefoged, P. and Whitteridge, D., Expiratory pressures and air flow during speech, Br. Med. J. 18, 1837–1843, 1960. Sears, T.A., Breathing: A sensori-motor act, Sci. Basis Med. Annu. Rev., 128–147, 1971. Roussos, C.S. and Macklem, P.T., Diaphragmatic fatigue in man, J. Appl. Physiol. 43, 189–197, 1977. Patakas, D., Louridas, G., Argyropoulou, P. and Stavropoulos, C., Respiratory drive in patients with chronic obstructive pulmonary disease, Respiration 36, 194–200, 1999. Cohen, E., Murphy, K., Adams, L., Guz, A. and Benchetrit, G., Is voluntary control of breathing impaired in patients with chronic obstructive pulmonary disease? Clin. Sci. (Lond.), 88, 453–461, 1995. Gorman, R.B., McKenzie, D.K. and Gandevia, S.C., Task failure, breathing discomfort and CO2 accumulation without fatigue during inspiratory resistive loading in humans, Respir. Physiol. 115, 273–286, 1999. Gandevia, S.C., Allen, G.M., Butler, J.E., Gorman, R.B. and McKenzie, D.K., Human respiratory muscles, sensations, reflexes and fatiguability, Clin. Exp. Pharmacol. Physiol. 25, 757–763, 1998. Topeli, A., Laghi, F. and Tobin, M.J., The voluntary drive to breathe is not decreased in hypercapnic patients with severe COPD, J. Eur. Respir. 18, 53–60, 2001. Gorini, M., Spinelli, A., Ginanni, R., Duranti, R., Gigliotti, F. and Scano, G., Neural respiratory drive and neuromuscular coupling in patients with Chronic Obstructive Pulmonary Disease (COPD), Chest 98, 1179–1186, 1990.
12. 13. 14. 15. 16. 17. 18. 19. 20.
21.
22. 23. 24. 25.
26.
27.
28.
29.
30.
Suprapontine Control of Breathing 31.
32.
33. 34. 35. 36. 37. 38.
39.
40.
41.
42. 43.
44.
45.
46.
93
Munschauer, F.E., Mador, M.J., Ahuja, A. and Jacobs, L., Selective paralysis of voluntary but not limbically influenced automatic respiration, Arch. Neurol. 48, 1190–1192, 1991. Heywood, P., Murphy, K., Corfield, D.R., Morrell, M.J., Howard, R.S. and Guz, A., Control of breathing in man, insights from the locked-in syndrome, Respir. Physiol. 106, 13–20, 1996. Foerster, O., Motorische Felden und Bahen, in Handbook der Neurologie. 6, Bumke, O., Foerster, O., eds., Berlin, Springer, pp. 50–51, 1936. Merton, P.A. and Morton, H.B., Stimulation of the cerebral cortex in the intact human subject, Nature 285, 227, 1980. Gandevia, S.C. and Rothwell, J.C., Activation of the human diaphragm from the motor cortex, J. Physiol. 384, 109–118, 1987. Barker, A.T., Jalinous, R. and Freeston, I.L., Non-invasive magnetic stimulation of human motor cortex, Lancet 1, 1106–1107, 1985. Maskill, D., Murphy, K., Mier, A., Owen, M. and Guz, A., Motor cortical representation of the diaphragm in man, J. Physiol. 443, 105–121, 1991. Fink, G.R., Corfield, D.R., Murphy, K., Kobayashi, I., Dettmers, C., Adams, L., Frackowiak, R.S. and Guz, A., Human cerebral activity with increasing inspiratory force: A study using positron emission tomography, J. Appl. Physiol. 81, 1295–1305, 1996. Colebatch, J.G., Adams, L., Murphy, K., Martin, A.J., Lammertsma, A.A., Tochon-Danguy, H.J., Clark, J.C., Friston, K.J. and Guz, A., Regional cerebral blood flow during volitional breathing in man, J. Physiol. 443, 91–103, 1991. Ramsay, S.C., Adams, L., Murphy, K., Corfield, D.R., Grootoonk, S., Bailey, D.L., Frackowiak, R.S. and Guz, A., Regional cerebral blood flow during volitional expiration in man: A comparison with volitional inspiration, J. Physiol. 461, 85–101, 1993. Evans, K.C., Shea, S.A. and Saykin, A.J., Functional MRI localisation of central nervous system regions associated with volitional inspiration in humans, J. Physiol. 520 Pt 2, 383–392, 1999. Smejkal, V., Druga, R. and Tintera, J., Brain activation during volitional control of breathing, Physiol. Res. 49, 659–663, 2000. McKay, L.C., Evans, K.C., Frackowiak, R.S. and Corfield, D.R., Neural correlates of voluntary breathing in humans and J. Appl. Physiol. 95, 1170–1178, 2003. Plum, F. and Leigh, R.J., Abnormalities of central mechanisms, in Regulation of Breathing Part II, Hornbein, T., ed., New York, Marcel Dekker, Inc., pp. 989–1067, 1981. Urban, P.P., Morgenstern, M., Brause, K., Wicht, S., Vukurevic, G., Kessler, S. and Stoeter, P., Distribution and course of cortico-respiratory projections for voluntary activation in man, a transcranial magnetic stimulation study in healthy subjects and patients with cerebral ischemia, J. Neurol. 249, 735–744, 2002. Boiten, F.A., Frijda, N.H. and Wientjes, C.J., Emotions and respiratory patterns, review and critical analysis, Int. J. Psychophysiol. 17, 103–128, 1994.
94
Moosavi et al.
47.
Cherniack, N.S., Lavietes, M.H., Tiersky, L. and Natelson, B.H., Respiratory sensations may be controlling elements on ventilation but can be affected by personality traits and state changes, in Respiration and Emotions, Haruki, Y., Homma, I., Umezawa, A. and Masaoka, Y., eds., Tokyo, Springer-Verlag, pp. 11–20, 2001. Banzett, R. and Lansing, R., Respiratory sensations arising from chemoreceptors and pulmonary receptors, air hunger and lung volume, in Respiratory Sensation, Vol. 90, Adams, L. and Guz, A., eds., New York, Marcel Dekker, pp. 155–180, 1996. Benussi, V., Die Atmungssymptome der Luge, Archiv. fur die Gesamte. Psychologie 31, 244–273, 1914. Feleky, A., The influence of the emotions on respiration, J. Exp. Psych. 1, 218–241, 1916. Dudley, D.L., Holmes, T.H., Martin, C.J. and Ripley, H.S., Changes in respiration associated with hypnotically induced emotion, pain and exercise, Psychosom. Med. 26, 46–57, 1964. Sebastiani, L., Simoni, A., Gemignani, A., Ghelarducci, B. and Santarcangelo, E.L., Human hypnosis, autonomic and electroencephalographic correlates of a guided multimodal cognitive-emotional imagery, Neurosci. Lett. 338, 41–44, 2003. May, J.R. and Johnson, H.J., Physiological activity to internally elicited arousal and inhibitory thoughts, J. Abnorm. Psychol. 82, 239–245, 1973. Ritz, T., Alatupa, S., Thons, M. and Dahme, B., Effects of affective picture viewing and imagery on respiratory resistance in nonasthmatic individuals, Psychophysiology 39, 86–94, 2002. Ax, A.F., The physiological differentiation between fear and anger in humans, Psychosom. Med. 15, 433–442, 1953. Suess, W.M., Alexander, A.B., Smith, D.D., Sweeney, H.W. and Marion, R.J., The effects of psychological stress on respiration, a preliminary study of anxiety and hyperventilation, Psychophysiology 17, 535–540, 1980. Heim, E., Knapp, P.H., Vachon, L., Globus, G.G. and Nemetz, S., Emotion, breathing and speech, J. Psychosom. Res. 12, 261–274, 1968. Hesson, K., Hill, T. and Bakal, D., Variability in breathing patterns during latent labor, a pilot study, J. Nurse Midwifery 42, 99–103, 1997. Boiten, F.A., The effects of emotional behavior on components of the respiratory cycle, Biol. Psychol. 49, 29–51, 1998. Takase, H. and Haruki, Y., Coordination of breathing between ribcage and abdomen in emotional arousal, in Respiration and Emotions, Haruki, Y., Homma, I., Umezawa, A. and Masaoka, Y., eds., Tokyo, Springer-Verlag, pp. 75–86, 2001. Mador, M.J. and Tobin, M.J., Effect of alterations in mental activity on the breathing pattern in healthy subjects, Am. Rev. Respir. Dis. 144, 481–487, 1991. Santibanez, G. and Bloch, S., A qualitative analysis of emotional effector patterns and their feedback, Pavlov. J. Biol. Sci. 21, 108–116, 1986. Grossman, P. and Wientjes, C.J., How breathing adjusts to mental and physical demands, in Respiration and Emotions, Haruki, Y., Homma, I., Umezawa, A. and Masaoka, Y., eds., Tokyo, Springer-Verlag, pp. 43–54, 2001.
48.
49. 50. 51.
52.
53. 54.
55. 56.
57. 58. 59. 60.
61. 62. 63.
Suprapontine Control of Breathing 64. 65. 66.
67. 68.
69.
70.
71. 72. 73. 74. 75. 76.
77. 78.
79. 80.
81.
82.
95
Paterson, A.S., The depth and rate of respiration in normal and psychotic subjects, J. Neurol. Psychopathol. 14, 322–331, 1934. Nixon, P.G. and Freeman, L.J., The think test, a further technique to elicit hyperventilation, J. R. Soc. Med. 81, 277–279, 1988. Yeragani, V.K., Radhakrishna, R.K., Tancer, M. and Uhde, T., Nonlinear measures of respiration, respiratory irregularity and increased chaos of respiration in patients with panic disorder, Neuropsychobiology 46, 111–120, 2002. Wilhelm, F.H., Trabert, W. and Roth, W.T., Characteristics of sighing in panic disorder, Biol. Psychiatry 49, 606–614, 2001. Martinez, J.M., Kent, J.M., Coplan, J.D., Browne, S.T., Papp, L.A., Sullivan, G.M., Kleber, M., Perepletchikova, F., Fyer, A.J., Klein, D.F., Gorman, J.M., Respiratory variability in panic disorder, Depress. Anxiety 14, 232–237, 2001. Sinha, S., Papp, L.A. and Gorman, J.M., How study of respiratory physiology aided our understanding of abnormal brain function in panic disorder, J. Affect. Disord. 61, 191–200, 2000. Filippelli, M., Pellegrino, R., Iandelli, I., Misuri, G., Rodarte, J.R., Durante, R., Brusasco, V. and Scano, G., Respiratory dynamics during laughter, J. Appl. Physiol. 90, 1441–1446, 2001. Hess, D.R. and Bigatello, L.M., Lung recruitment, the role of recruitment maneuvers, Respir. Care 47, 308–317, discussion 317–318, 2002. Grossman, P., Respiration, stress, and cardiovascular function, Psychophysiology 20, 284–300, 1983. Alpher, V.S. and Blanton, R.L., Motivational processes and behavioral inhibition in breath holding, J. Psychol. 125, 71–81, 1991. James, W., What is an emotion? Mind 19, 188–205, 1884. Lehmann, H.E. and Yawning, A homeostatic reflex and its psychological significance, Bull. Menninger Clin. 43, 123–126, 1979. Platek, S.M., Critton, S.R., Myers, T.E. and Gallup, G.G., Contagious yawning, the role of self-awareness and mental state attribution, Brain Res. Cogn. Brain Res. 17, 223–227, 2003. Muchnik, S., Finkielman, S., Semeniuk, G., de Aguirre, M.I. and [Yawning], Medicina Buenos Aires, 63, 229–232, 2003. Phillips, M.L., Drevets, W.C., Rauch, S.L. and Lane, R., Neurobiology of emotion perception I, The neural basis of normal emotion perception, Biol. Psychiatry 54, 504–514, 2003. Bauer, G., Gerstenbrand, F. and Hengl, W., Involuntary motor phenomena in the locked-in syndrome, J. Neurol. 223, 191–198, 1980. Cherniack, N.S., Respiratory sensation as a respiratory controller, in Respiratory Sensation, Vol. 90, Adams, L. and Guz, A., eds., New York, Marcel Dekker, pp. 213–225, 1996. Shea, S.A., Banzett, R.B. and Lansing, R.W., Respiratory sensations and their role in the control of breathing, in Regulation of Breathing, Vol. 79, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 923–957, 1996. Simon, P.M., Schwartzstein, R.M., Weiss, J.W., Lahive, K., Fencl, V., Teghtsoonian, M. and Weinberger, S.E., Distinguishable sensations of
96
83.
84.
85.
86.
87.
88.
89. 90.
91.
92.
93.
94.
95.
96.
97.
Moosavi et al. breathlessness induced in normal volunteers, Am. Rev. Respir. Dis. 140, 1021–1027, 1989. Demediuk, B.H., Manning, H., Lilly, J., Fencl, V., Weinberger, S.E., Weiss, J.W. and Schwartzstein, R.M., Dissociation between dyspnea and respiratory effort, Am. Rev. Respir. Dis. 146, 1222–1225, 1992. Simon, P.M., Schwartzstein, R.M., Weiss, J.W., Fencl, V., Teghtsoonian, M. and Weinberger, S.E., Distinguishable types of dyspnea in patients with shortness of breath, Am. Rev. Respir. Dis. 142, 1009–1014, 1990. Elliott, M.W., Adams, L., Cockcroft, A., MacRae, K.D., Murphy, K. and Guz, A., The language of breathlessness, use of verbal descriptors by patients with cardiopulmonary disease, Am. Rev. Respir. Dis. 144, 826–832, 1991. Moosavi, S.H., Topulos, G.P., Hafer, A., Lansing, R.W., Adams, L., Brown, R. and Banzett, R.B., Acute partial paralysis alters perceptions of air hunger, work and effort at constant P(CO(2)) and V(E), Respir. Physiol. 122, 45–60, 2000. Lansing, R.W., Im, B.S., Thwing, J.I., Legedza, A.T. and Banzett, R.B., The perception of respiratory work and effort can be independent of the perception of air hunger, Am. J. Respir. Crit. Care Med. 162, 1690–1696, 2000. Killian, K.J., Gandevia, S.C., Summers, E. and Campbell, E.J., Effect of increased lung volume on perception of breathlessness, effort, and tension, J. Appl. Physiol. 57, 686–691, 1984. Gandevia, S.C., Killian, K.J. and Campbell, E.J., The effect of respiratory muscle fatigue on respiratory sensations, Clin. Sci. (Lond.), 60, 463–466, 1981. Binks, A.P., Moosavi, S.H., Banzett, R.B. and Schwartzstein, R.M., Tightness sensation of asthma does not arise from the work of breathing, Am. J. Respir. Crit. Care Med. 165, 78–82, 2002. Moy, M.L., Woodrow Weiss, J., Sparrow, D., Israel, E. and Schwartzstein, R.M., Quality of dyspnea in bronchoconstriction differs from external resistive loads, Am. J. Respir. Crit. Care Med. 162, 451–455, 2000. Cherniack, N.S., Potential role of optimization in alveolar hypoventilation and respiratory instability, in Neurobiology of the Control of Breathing, Euler, C.V. and Lagercrantz, H., eds., New York, Raven Press, pp. 45–50, 1986. Wilson, R.C. and Jones, P.W., Differentiation between the intensity of breathlessness and the distress it evokes in normal subjects during exercise, Clin. Sci. (Lond.) 80, 65–70, 1991. Carrieri-Kohlman, V., Gormley, J.M., Douglas, M.K., Paul, S.M. and Stulbarg, M.S., Differentiation between dyspnea and its affective components, West J. Nurs. Res. 18, 626–642, 1996. Rainville, P., Feine, J.S., Bushnell, M.C. and Duncan, G.H., A psychophysical comparison of sensory and affective responses to four modalities of experimental pain, Somatosens. Mot. Res. 9, 265–277, 1992. Banzett, R.B. and Moosavi, S.H., Dyspnea and pain, similarities and contrasts between two very unpleasant sensations, Am. Pain Soc. Bull. 11, 1 (and 6–8), 2001. Murphy, K., Mier, A., Adams, L. and Guz, A., Putative cerebral cortical involvement in the ventilatory response to inhaled CO2 in conscious man, J. Physiol. 420, 1–18, 1990.
Suprapontine Control of Breathing 98.
99.
100.
101. 102. 103.
104.
105.
106.
107.
108.
109.
110.
111.
97
Shea, S.A., Harty, H.R. and Banzett, R.B., Self-control of level of mechanical ventilation to minimize CO2 induced air hunger, Respir. Physiol. 103, 113–125, 1996. Corfield, D.R., Fink, G.R., Ramsay, S.C., Murphy, K., Harty, H.R., Watson, J.D., Adams, L., Frackowiak, R.S. and Guz, A., Evidence for limbic system activation during CO2-stimulated breathing in man, J. Physiol. 488 (Pt 1), 77–84, 1995. Gozal, D., Hathout, G.M., Kirlew, K.A., Tang, H., Woo, M.S., Zhang, J., Lufkin, R.B. and Harper, R.M., Localization of putative neural respiratory regions in the human by functional magnetic resonance imaging, J. Appl. Physiol. 76, 2076–2083, 1994. Brashear, R.E., Hyperventilation syndrome, Lung 161, 257–273, 1983. Lewis, R.A. and Howell, J.B., Definition of the hyperventilation syndrome, Bull. Eur. Physiopathol. Respir. 22, 201–205, 1986. Van Diest, I., Proot, P., Van De Woestijne, K.P., Han, J.N., Devriese, S., Winters, W. and Van Den Bergh, O., Critical conditions for hyperventilation responses, The role of autonomic response propositions during emotional imagery, Behav. Modif. 25, 621–639, 2001. Jack, S., Wilkinson, M. and Warburton, C.J., Behavioral and physiological factors affecting breathing pattern and ventilatory control in patients with idiopathic hyperventilation, in Respiration and Emotions, Haruki, Y., Homma, I., Umezawa, A. and Masaoka, Y., eds., Tokyo, Springer-Verlag, pp. 87–98, 2001. Kikuchi, Y., Okabe, S., Tamura, G., Hida, W., Homma, M., Shirato, K. and Takishima, T., Chemosensitivity and perception of dyspnea in patients with a history of near-fatal asthma, N. Engl. J. Med. 330, 1329–1334, 1994. Lanini, B., Gigliotti, F., Coli, C., Bianchi, R., Pizzi, A., Romagnoli, I., Grazzini, M., Stendardi, L. and Scano, G., Dissociation between respiratory effort and dyspnoea in a subset of patients with stroke, Clin. Sci. (Lond.), 103, 467–473, 2002. Allen, G.M., Hickie, I., Gandevia, S.C. and McKenzie, D.K., Impaired voluntary drive to breathe, a possible link between depression and unexplained ventilatory failure in asthmatic patients, Thorax 49, 881–884, 1994. Peiffer, C., Poline, J.B., Thivard, L., Aubier, M. and Samson, Y., Neural substrates for the perception of acutely induced dyspnea, Am. J. Respir. Crit. Care Med. 163, 951–957, 2001. Banzett, R.B., Mulnier, H.E., Murphy, K., Rosen, S.D., Wise, R.J. and Adams, L., Breathlessness in humans activates insular cortex, Neuroreport 11, 2117–2120, 2000. Brannan, S., Liotti, M., Egan, G., Shade, R., Madden, L., Robillard, R., Abplanalp, B., Stofer, K. and Denton, D., Neuroimaging of cerebral activations and deactivations associated with hypercapnia and hunger for air, Proc. Natl. Acad. Sci. USA 98, 2029–2034, 2001. Evans, K.C., Banzett, R.B., Adams, L., McKay, L., Frackowiak, R.S. and Corfield, D.R., BOLD fMRI identifies limbic, paralimbic, and cerebellar activation during air hunger, J. Neurophysiol. 88, 1500–1511, 2002.
98
Moosavi et al.
112. Augustine, J.R., Circuitry and functional aspects of the insular lobe in primates including humans, Brain Res. Rev. 22, 229–244, 1996. 113. Gaytan, S.P. and Pasaro, R., Connections of the rostral ventral respiratory neuronal cell group, an anterograde and retrograde tracing study in the rat, Brain Res. Bull. 47, 625–642, 1998. 114. Hanamori, T., Kunitake, T., Kato, K. and Kannan, H., Responses of neurons in the insular cortex to gustatory, visceral, and nociceptive stimuli in rats, J. Neurophysiol. 79, 2535–2545, 1998. 115. Hanamori, T., Kunitake, T., Kato, K. and Kannan, H., Neurons in the posterior insular cortex are responsive to gustatory stimulation of the pharyngolarynx, baroreceptor and chemoreceptor stimulation, and tail pinch in rats, Brain Res. 785, 97–106, 1998. 116. Kinomura, S., Kawashima, R., Yamada, K., Ono, S., Itoh, M., Yoshioka, S., Yamaguchi, T., Matsui, H., Miyazawa, H. and Itoh, H., et al., Functional anatomy of taste perception in the human brain studied with positron emission tomography, Brain Res. 659, 263–266, 1994. 117. Miller, A.D., Rowley, H.A., Roberts, T.P. and Kucharczyk, J., Human cortical activity during vestibular- and drug-induced nausea detected using MSI, Ann. N.Y. Acad. Sci. 781, 670–672, 1996. 118. Casey, K.L., Forebrain mechanisms of nociception and pain, analysis through imaging, Proc. Natl. Acad. Sci. USA 96, 7668–7674, 1999. 119. Banzett, R., Lansing, R., Evans, K. and Shea, S., Stimulus-response characteristics of CO2-induced air hunger in normal subjects, Respir. Physiol. 103, 19–31, 1996. 120. Chen, Z., Eldridge, F.L. and Wagner, P.G., Respiratory-associated thalamic activity is related to level of respiratory drive, Respir. Physiol. 90, 99–113, 1992. 121. Chen, Z., Eldridge, F.L. and Wagner, P.G., Respiratory-associated rhythmic firing of midbrain neurones in cats, relation to level of respiratory drive, J. Physiol. 437, 305–325, 1991. 122. Banzett, R.B., Lansing, R.W., Brown, R., Topulos, G.P., Yager, D., Steele, S.M., Londono, B., Loring, S.H., Reid, M.B. and Adams, L., et al., Air hunger from increased PCO2 persists after complete neuromuscular block in humans, Respir. Physiol. 81, 1–17, 1990. 123. Gandevia, S.C., Killian, K., McKenzie, D.K., Crawford, M., Allen, G.M., Gorman, R.B. and Hales, J.P., Respiratory sensations, cardiovascular control, kinaesthesia and transcranial stimulation during paralysis in humans, J. Physiol. (Lond.) 470, pp. 85–107, 1993. 124. Flume, P., Eldridge, F., Edwards, L. and Houser, L., The Fowler breathholding study revisited, continuous rating of respiratory sensation, Respir. Physiol. 95, 53–66, 1994. 125. Flume, P.A., Eldridge, F.L., Edwards, L.J. and Mattison, L., Relief of the air hunger of breathholding, A role for pulmonary stretch receptors, Respir. Physiol. 103, 221–232, 1996. 126. Tobin, M.J., Perez, W., Guenther, S.M., D’Alonzo, G. and Dantzker, D.R., Breathing pattern and metabolic behavior during anticipation of exercise, J. Appl. Physiol. 60, 1306–1312, 1986.
Suprapontine Control of Breathing
99
127. Krogh, A. and Lindhard, J., The regulation of respiration and circulation during the initial stages of muscular work, J. Physiol. 47, 112–136, 1913. 128. Masaoka. Y. and Homma, I., The effect of anticipatory anxiety on breathing and metabolism in humans, Respir. Physiol. 128, 171–177, 2001. 129. Poole, R., Goetzinger, C.P. and Rousey, C.L., A study of the effects of auditory stimuli on respiration, Acta Otolaryngol. 61, 143–152, 1966. 130. Skaggs, E.B., Changes in pulse, breathing and steadiness under conditions of startledness and excited expectancy, J. Comp. Psychol. 6, 303–318, 1926. 131. Harver, A. and Kotses, H., Effects of auditory stimulation on respiration, Psychophysiology 24, 26–34, 1987. 132. Gautier, H., Respiratory and heart rate responses to auditory stimulations, Physiol. Behav. 8, 327–332, 1972. 133. Shea, S.A., Walter, J., Pelley, C., Murphy, K. and Guz, A., The effect of visual and auditory stimuli upon resting ventilation in man, Respir. Physiol. 68, 345–357, 1987. 134. Harrer, G., Somatische aspekte des musikerlebeus, Med. Monatspiegel. 6, 124–127, 1970. 135. Haas, F., Distenfeld, S. and Axen, K., Effects of perceived musical rhythm on respiratory pattern, J. Appl. Physiol. 61, 1185–1191, 1986. 136. Haun, M., Mainous, R.O. and Looney, S.W., Effect of music on anxiety of women awaiting breast biopsy, Behav. Med. 27, 127–132, 2001. 137. Wong, H.L., Lopez-Nahas, V. and Molassiotis, A., Effects of music therapy on anxiety in ventilator-dependent patients, Heart Lung 30, 376–387, 2001. 138. Blood, A.J., Zatorre, R.J., Bermudez, P. and Evans, A.C., Emotional responses to pleasant and unpleasant music correlate with activity in paralimbic brain regions, Nat. Neurosci. 2, 382–387, 1999. 139. Fink, B.R., The influence of cerebral activity in wakefulness on regulation of breathing, J. Appl. Physiol. 16, 15–20, 1961. 140. Morrell, M.J., Shea, S.A., Adams, L. and Guz, A., Effects of inspiratory support upon breathing in humans during wakefulness and sleep, Respir. Physiol. 93, 57–70, 1993. 141. Datta, A.K., Shea, S.A., Horner, R.L. and Guz, A., The influence of induced hypocapnia and sleep on the endogenous respiratory rhythm in humans, J. Physiol. 440, 17–33, 1991. 142. Skatrud, J.B. and Dempsey, J.A., Interaction of sleep state and chemical stimuli in sustaining rhythmic ventilation, J. Appl. Physiol. 55, 813–822, 1983. 143. Fink, B.R., Hanks, E.C., Ngai, S.H. and Papper, E.M., Central regulation of respiration during anesthesia and wakefulness, Ann. N.Y. Acad. Sci. 109, 892–900, 1963. 144. Orem, J., The nature of the wakefulness stimulus for breathing, in Sleep and Respiration, Issa, F.G., Suratt, P.M. and Remmers, J.E., eds., New York, Wiley-Liss, pp. 23–31, 1990. 145. Hugelin, A. and Cohen, M.I., The reticular activating system and respiratory regulation in the cat, Ann. N.Y. Acad. Sci. 109, 586–603, 1963. 146. Saper, C.B., Chou, T.C. and Scammell, T.E., The sleep switch, hypothalamic control of sleep and wakefulness, Trends Neurosci. 24, 726–731, 2001.
100
Moosavi et al.
147. Gothe, B., Altose, M.D., Goldman, M.D. and Cherniack, N.S., Effect of quiet sleep on resting and CO2-stimulated breathing in humans, J. Appl. Physiol. 50, 724–730, 1981. 148. Orem, J., Montplaisir, J. and Dement, W.C., Changes in the activity of respiratory neurons during sleep, Brain Res. 82, 309–315, 1974. 149. Phillipson, E.A. and Bowes, G., Control of breathing during sleep, in Handbook of Physiology, The Respiratory System II, Chapter 19, Cherniack, N.S. and Widdicombe, J.G., eds., Bethesda, MD, American Physiological Society, pp. 649–689, 1986. 150. Shea, S.A., Life without ventilatory chemosensitivity, Respir. Physiol. 110, 199–210, 1997. 151. Kaada, B.R., Cingulate, posterior orbital, anterior insula and temporal pole cortex, in Handbook of Physiology, Magoun, H.W., ed., Washington, DC, American Physiological Society, pp. 1345–1372, 1960. 152. Nelson, D.A. and Ray, C.D., Respiratory arrest from seizure discharges in limbic system, Report of cases, Arch. Neurol. 19, 199–207, 1968. 153. Brown, H.W. and Plum, F., The neurological basis of Cheyne Stokes respiration, Am. J. Med. 30, 849–860, 1961. 154. Heyman, A.R., Birchfield, R.I. and Sieker, H.O., Effects of bilateral cerebral infarction on respiratory center sensitivity, Neurology 8, 694–700, 1958. 155. Tenney, S.M. and Ou, L.C., Ventilatory response of decorticate and decerebrate cats to hypoxia and CO2, Respir. Physiol. 29, 81–92, 1977. 156. Reis, D.J. and McHugh, P.R., Hypoxia as a cause of bradycardia during amygdala stimulation in monkey, Am. J. Physiol. 214, 601–610, 1968. 157. Feldman, J.L., Neurophysiology of breathing in mammals, in Handbook of Physiology The Nervous System IV, Bloom, F.E., ed., Bethesda, American Physiological Society, pp. 463–524, 1986. 158. Shik, M.L. and Orlovsky, G.N., Neurophysiology of locomotor automatism, Physiol. Rev. 56, 465–501, 1976. 159. Phillipson, E.A., McClean, P.A., Sullivan, C.E. and Zamel, N., Interaction of metabolic and behavioral respiratory control during hypercapnia and speech, Am. Rev. Respir. Dis. 117, 903–909, 1978. 160. Aminoff, M.J. and Sears, T.A., Spinal integration of segmental, cortical and breathing inputs to thoracic respiratory motoneurones, J. Physiol. 215, 557–575, 1971. 161. Davis, J.N. and Plum, F., Separation of descending spinal pathways to respiratory motoneurons, Exp. Neurol. 34, 78–94, 1972. 162. Orem, J. and Netick, A., Behavioral control of breathing in the cat, Brain Res. 366, 238–253, 1986. 163. Gallego, J. and Gaultier, C., [Respiratory behavior] Rev. Mal. Respir. 17, 41–49, 2000. 164. Mador, M.J. and Acevedo, F.A., Effect of respiratory muscle fatigue on subsequent exercise performance, J. Appl. Physiol. 70, 2059–2065, 1991. 165. Roussos, C., Fixley, M., Gross, D. and Macklem, P.T., Fatigue of inspiratory muscles and their synergic behavior, J. Appl. Physiol. 46, 897–904, 1979.
Suprapontine Control of Breathing
101
166. Ward, S.A., Macias, D. and Whipp, B.J., Is breath-hold time an objective index of exertional dyspnoea in humans? Eur. J. Appl. Physiol. 85, 272–279, 2001. 167. Murry, T., Nelson, E.J. and Swenson, E.W., Speech intelligibility during exercise at normal and increased atmospheric pressures, Aerosp. Med. 43, 887–890, 1972. 168. Bjurstrom, R.L. and Schoene, R.B., Control of ventilation in elite synchronized swimmers, J. Appl. Physiol. 63, 1019–1024, 1987. 169. Dejours, P., Control of respiration in muscular exercise, in Handbook of Physiology, Sec. 3, Respiration, Vol. 1, Chapter 25, Fenn, W.O. and Rahn, H., eds., Washington DC, American Physiological Society, pp. 631–648, 1964. 170. Bunn, J.C. and Mead, J., Control of ventilation during speech, J. Appl. Physiol. 31, 870–872, 1971. 171. Doust, J.H. and Patrick, J.M., The limitation of exercise ventilation during speech, Respir. Physiol. 46, 137–147, 1981. 172. Meckel, Y., Rotstein, A. and Inbar, O., The effects of speech production on physiologic responses during submaximal exercise, Med. Sci. Sports Exerc. 34, 1337–1343, 2002. 173. Shea, S.A., Hoit, J.D. and Banzett, R.B., Competition between gas exchange and speech production in ventilated subjects, Biol. Psychol. 49, 9–27, 1998. 174. Lee, L., Loudon, R.G., Jacobson, B.H. and Stuebing, R., Speech breathing in patients with lung disease, Am. Rev. Respir. Dis. 147, 1199–1206, 1993. 175. Martin, J.G., Shore, S.A. and Engel, L.A., Mechanical load and inspiratory muscle action during induced asthma, Am. Rev. Respir. Dis. 128, 455–460, 1983. 176. Sharp, J.T., Drutz, W.S., Moisan, T., Foster, J. and Machnach, W., Postural relief of dyspnea in severe chronic obstructive pulmonary disease, Am. Rev. Respir. Dis. 122, 201–211, 1980. 177. Ugalde, V., Breslin, E.H., Walsh, S.A., Bonekat, H.W., Abresch, R.T. and Carter, G.T., Pursed lips breathing improves ventilation in myotonic muscular dystrophy, Arch. Phys. Med. Rehabil. 81, 472–478, 2000. 178. Spahija, J.A. and Grassino, A., Effects of pursed-lips breathing and expiratory resistive loading in healthy subjects, J. Appl. Physiol. 80, 1772–1784, 1996. 179. Bai, C.X., [Application of pursed lips breathing to chronic obstructive pulmonary disease patients with respiratory insufficiency] Zhonghua Jie He He Hu Xi Za Zhi 14, 283–284, 319, 1991. 180. Banzett, R.B., Topulos, G.P., Leith, D.E. and Nations, C.S., Bracing arms increases the capacity for sustained hyperpnea, Am. Rev. Respir. Dis. 138, 106–109, 1988. 181. Prandi, E., Couture, J. and Bellemare, F., In normal subjects bracing impairs the function of the inspiratory muscles, Eur. Respir. J. 13, 1078–1085, 1999. 182. Gallego, J., Ankaoua, J., Camus, J.F. and Jacquemin, C., Synchronization and knowledge of results in a ventilatory motor task., Percept. Mot. Skills 63, 3–10, 1986.
102
Moosavi et al.
183. Johnston, R. and Lee, K., Myofeedback, a new method of teaching breathing exercises in emphysematous patients, Phys. Ther. 56, 826–831, 1976. 184. Donner, C.F. and Howard, P., Pulmonary rehabilitation in chronic obstructive pulmonary disease (COPD) with recommendations for its use, Report of the European Respiratory Society Rehabilitation and Chronic Care Scientific Group (S.E.P.C.R. Rehabilitation Working Group) Eur. Respir. J. 5, 266–275, 1992. 185. van Doorn, P., Folgering, H. and Colla, P., Control of the end-tidal PCO2 in the hyperventilation syndrome, effects of biofeedback and breathing instructions compared, Bull. Eur. Physiopathol. Respir. 18, 829–836, 1982. 186. Cowley, D.S. and Roy-Byrne, P.P., Hyperventilation and panic disorder, Am. J. Med. 83, 929–937, 1987. 187. Somjen, G., The missing error signal—regulation beyond negative feedback, News in Physiological Sciences 7, 184–185, 1992. 188. Shik, L.L., [Regulation of respiration during muscular exertion] Nauchnye Doki. Vyss. Shkoly. Biol. Nauki. 18–29, 1985. 189. Martin, P.A. and Mitchell, G.S., Long-term modulation of the exercise ventilatory response in goats, J. Physiol. 470, 601–617, 1993. 190. Helbling, D., Boutellier, U. and Spengler, C.M., Modulation of the ventilatory increase at the onset of exercise in humans, Respir. Physiol. 109, 219–229, 1997. 191. Turner, D.L. and Sumners, D.P., Associative conditioning of the exercise ventilatory response in humans, Respir. Physiol. Neurobiol. 132 159–168, 2002. 192. Moosavi, S.H., Guz, A. and Adams, L., Repeated exercise paired with imperceptible dead space loading does not alter VE of subsequent exercise in humans, J. Appl. Physiol. 92, 1159–1168, 2002. 193. Bykov, K., The Cerebral Cortex and the Internal Organs, New York, Chemical, 1957. 194. Gallego, J. and Perruchet, P., Classical conditioning of ventilatory responses in humans, J. Appl. Physiol. 70, 676–682, 1991.
4 Measurement of Drug Effects on Ventilatory Control
DENHAM S. WARD University of Rochester School of Medicine and Dentistry Rochester, New York
I.
Introduction
Drug effect studies are generally performed either to determine the effects (or side effects) of a clinically used drug, or to use the drug as a pharmacological probe to learn more about the functioning of ventilatory control mechanisms. An example of the former might be the comparison of the ventilatory depressing effects of different opioids [1] and of the latter, the use of a receptor antagonist to determine if an endogenous neurotransmitter is involved in ventilatory control [2]. Many techniques have been used to study and quantify drug effects on the control of breathing, but few standard methods for clinical testing have evolved. This chapter will primarily review methodology used to characterize drug effects on ventilatory control in humans in order to obtain useful clinical or physiological information. The other chapters in this book provide a wide range of the many different techniques that have been used, both in human and in invasive animal studies. Few drugs are studied for their positive effects on ventilatory control. Examples might be the use of almitrine [3,4] or doxapram [5] as ventilatory 103
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stimulants or the effects of naloxone [6] or flumazenil [7] in reversing the respiratory depressing effects of opioids or benzodiazepines. More often drug effect studies on the control of breathing are investigations of the ventilatory side effects of drugs used, for example, to provide pain relief. When studying the effects of drugs on the ventilatory control system, both the drug characteristics (pharmacokinetic and pharmacodynamic) and the difficulties present in making measurements on the ventilatory controller must be taken into account. This can present experimental problems quite different from those seen when experiments are designed to test a physiological hypothesis. However, most tests for drug effects use techniques originally designed as physiological experiments, often in animal models. There is no agreement if a particular test best predicts clinically important outcomes (e.g., post-operative respiratory complications and use in obstructive sleep apneic patients) or is best used to gain physiological information. Recurring themes in drug effect studies include how to handle the closed-loop nature of the respiratory controller and whether or not stimulated (e.g., with inhalation of CO2) or unstimulated (resting) measurements best reflect the drug effect [8,9]. If the clinical situation is studied, the feedback loops are left intact and no stimulation is applied. However, the well-known insensitivity of the controlled variables (outputs) to system parameter variation in a negative feedback control system often limits the observed effect of the drug, resulting in small changes in the measured variables [10]. This is illustrated in Figure 4.1 with a hypothetical drug causing general anesthesia and depression of the hypercapnic ventilatory response. If the drug effect is measured by the change in resting ventilation, then only a small effect is observed. Since the metabolic hyperbola is relatively flat around the normal resting values, there is a moderate increase in the PaCO2, but if the decrease in ventilation at a fixed hypercapnic level (e.g., PaCO2 ¼ 55 mm Hg) is measured, then there is a very large drug effect. When various maneuvers are used to open the feedback loops, the effect of the drug may be more apparent but the clinical interpretation may be called into question [9]. The classic investigation of the respiratory control system is to characterize the input–output relationship (static or dynamic) of all or part of the system and then determine how this relationship is modified by the administration of a drug. Invasive animal studies can focus on a single component of the system such as the carotid sinus nerve firing rate. However, the complete intact system has many inputs and outputs, as well as many internal feedback loops besides the dominant CO2-ventilation closed loop. These feedback loops present considerable difficulty in reliably estimating system performance when characteristics or parameters of the individual components are measured. In humans or in intact animal preparations, inputs to the control system can be divided into closed-loop
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Figure 4.1 Effect of a hypothetical anesthetic on measurements of the ventilatory control. This drug causes a large reduction in the hypercapnic response slope, a relatively small increase (shift to the right) of the zero ventilation intercept (2 mm Hg), and an elimination of the wakefulness drive to breath. This results in a relatively small decrease in the alveolar ventilation at rest (2 l/min); but because of the flatness of the metabolic hyperbola a larger increase in the resting CO2 (15 mm Hg). However, if the drug effect was quantified by the reduction of ventilation at a fixed CO2 (55 mm Hg has been commonly used—V55), a large decrease in ventilation is measured (30 l/min).
(chemoreflex as labeled by Dejours [11]) and open-loop (non-chemoreflex) [12]. The sensitivity of a chemoreflex (e.g., hypercapnic chemoreflex) is often characterized by a gain with units related to the change in output divided by the change in input (e.g., l min1 mm Hg1), but the fact that this gain is determined by all factors from input to output is important to remember. It is also tempting to consider the non-chemoreflex inputs as those setting the baseline or resting ventilation, and the chemoreflexes as those setting the gain of the closed loops; however, it is important to note that the non-chemoreflex inputs can modulate the chemoreflexes (see Ward and Karan [13] and Chapter 3, this volume, for recent reviews). Many of the non-chemoreflexes, including volitional inputs [14], have projections into or through the brainstem respiratory centers [15] and thus have the potential to alter the chemoreflex loop gain. For example, in nonrapid-eye movement (non-REM) sleep the hypoxic response is markedly reduced with only a small decrease in baseline ventilation [16,17]. Also, it has long been known that sleep and opioids have a strong interaction in
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depressing ventilation [18,19]. These interactions underscore the need to carefully design and perform experiments assessing drug effects on ventilatory control. An essential part of assessing drug effects is the parameterization of the respiratory response. This requires some type of model, statistical or parametric, that can be fitted to the respiratory data. Today, breathby-breath measurements are the norm and are used as the raw data for the models. Modeling in the control of breathing has had a long history [20] (see Khoo and Yamashiro [21] for a recent review); these models have played an integral part in the development and validation of methods to assess the ventilatory control system [22]. However, these models often require many parameters determined from several experiments. If the model parameters are to be estimated from the measured data of a single brief experiment, as is usually required when testing for a drug effect, then a greatly simplified model is needed. For the clinical assessment of drugs (and frequently used drug combinations), there might be expected some degree of standardization on the appropriate approach. However, this is not the case and there is a very wide range of published techniques. In the design of a study to ascertain a drug’s clinical effects, many factors need to be considered (Table 4.1) and the principles of good clinical trial design applied, whether the study is an outcome clinical trial or a laboratory study [23]. For example, the particular study design will have a major impact on the required size of the study. Thus, a laboratory investigating the effect of a drug on a particular chemoreflex, with a carefully selected, homogeneous subject population and with the drug levels tightly controlled, may require only a few subjects to demonstrate an effect. However, a clinical trial attempting to show a change in the post-operative respiratory morbidity with one analgesic regimen versus another may require a large number of subjects. II.
Measurement Techniques
Standard measurement techniques include airway gas concentrations (partial pressures) and flows, usually on a continuous basis and using measurement devices with very fast response times. These continuous signals are then digitized with an analog-to-digital converter and stored in a digital computer. For most breath-to-breath data calculations a sample rate of 50 Hz is sufficient. This digital data can then be processed to calculate a set of commonly utilized respiratory parameters, e.g., inspired and end-tidal (PET) O2 and CO2, tidal volume, inspiratory time, expiratory time, minute ventilation, etc. Both commercial and laboratory-designed hardware and software are commonly used for these purposes [24]. Although there are many technical difficulties in making these measurements
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Table 4.1 Example Characteristics of Studies to Determine the Ventilatory Effects of Drugs in Human Subjects (1)
Clinical study (a) Clinical setting (e.g., recovery room, post-operative, out-patient) (b) Patient population (e.g., age, sex, weight, co-existing diseases, concurrent medications) (c) Drug administration method (e.g., patient-controlled analgesia, spinal, oral) (d) Stimulation (e.g., none, hypoxia, hypercapnia, negative airway pressure) (e) Outcome measurement (e.g., hypoxia, ventilation, complications)
(2)
Laboratory study (a) Subject population (e.g., normal, age, sex) (b) Drug administration (e.g., target controlled infusion, measurement of plasma levels) (c) Stimulation (i) Closed loop (chemoreflex) (e.g., hypoxia, hypercapnia, acidosis; ramps vs. steps) (ii) Non-closed loop (e.g., airway load, exercise, pain, audiovisual) (iii) Combined chemoreflex and non-chemoreflex stimulation (d) Measurements (e.g., ventilation, tidal volume, frequency, abdominal/chest wall excursions, esophageal/airway pressure, genioglossal EMG) (e) Response model (e.g., steady-state vs. transient, mixed effect, population PK-PD)
and ensuring proper instrument calibration, the problems have been well worked out and there should be no issues in making accurate measurements in the laboratory [25]. Confirmation of the measured end-tidal values with arterial blood gas measurements is a good practice when a new protocol or new equipment is used. An accurate, motor-driven syringe pump providing a consistent sinusoidal volume waveform is also invaluable in the set-up and maintenance of a flow/volume measurement system. Several reviews are available on the technical aspects of these measurement techniques (including a still useful special section in Chest, 70, 109–195, 1976) [26–29]. Measurement of arterial oxygenation is often made with pulse oximetry rather than with measurement of end-tidal O2. Although the PETO2 can often differ from the arterial PO2 by several mm Hg, the measurement of saturation with pulse oximetry can also be inaccurate and has a time delay that can differ between rapidly increasing versus decreasing oxygen tensions [30]. Since oxygenation is critically important, monitoring of patients who are at risk for respiratory depression with a pulse oximeter has become standard, both in routine clinical practice and as an outcome in clinical trials studying respiratory depression [31–35].
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Wheatley et al. used timed-epoch (1 h) histograms of the continuous saturation to show differences in the patterns of desaturation between pre- and post-surgery [34]. Obviously, while severe prolonged hypoxemia is dangerous, less severe episodic hypoxemia (e.g., in the post-operative period) has not been clearly linked to adverse outcomes [36]. However, chronic episodic desaturations (e.g., in sleep apnea patients) are more clearly associated with changes in health status [37,38]. While much work has focused on the pharmacologic alteration of the chemoreflexes, upper airway obstruction is clinically a predominant drug effect in many patients, especially those with obstructive sleep apnea. Inductance plethysmography has been used for many years to measure the relative motions of the ribcage and diaphragm [39] and to prevent some of the alterations in respiratory pattern caused by airway instrumentation [40,41]. Determination of upper airway obstruction can be done using inductance plethysmography and analyzing the signals for asynchrony [42]. However, other drug effects may cause asynchrony between the rib cage and abdomen without any overt airway obstruction and caution must be taken in interpreting these signals [43–45]. In fact, alterations in the relative amount of respiratory drive to the diaphragm versus the intercostal muscles, as assessed by inductance plethysmography, is an important drug effect, particularly in infants [46]. Inductance plethysmography may also be useful in clinical studies where long-term respiratory monitoring is needed and thus direct measurement of ventilation via airway instrumentation is not practical [35]. For many studies, the subject can breath through either a facemask or mouthpiece. A facemask is preferred since it causes less disruption of the normal respiratory pattern and permits normal nasal breathing. However, when testing sedatives, it is important that a good mask seal is maintained even when the mouth muscles become slack.
III.
Quantification of Drug Pharmacodynamics
To fully characterize the effects of a drug on ventilatory control, it is necessary to construct a full dose (or even better, a plasma concentration) response curve. This curve can then be parameterized with standardized pharmacodynamic parameters (e.g., plasma concentration for a 50% maximum effect, EC50). Selecting the particular respiratory responses to parameterize as well as the particular equation to use are important decisions. Since respiratory depression is often being evaluated, it is common for a study to select a dose in the therapeutic range (e.g., for an opioid it could be a dose providing adequate analgesia) and then investigate the respiratory depression for that single dose. However, much more
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information about the relative effects of respiratory depressants is obtained if full drug concentration–response curves are investigated. For example, Mildh et al. [47] used computer-controlled infusions of fentanyl and alfentanil to set plasma levels. They then used measurements of resting ventilation, PaCO2 and arterial plasma drug levels to fit either a Hill (ventilation) or a linear (PaCO2) equation. This approach permitted them to compare potencies of the drugs for respiratory depression. Detailed modeling of both the pharmacokinetic and the pharmacodynamic effects of drugs can be done; this may allow a better understanding of the mechanisms of the differences between drugs. Bragg et al. used a population-based approach to model the pharmacodynamics and kinetics of morphine and fentanyl to determine how developmental age changes the respiratory depression of the drugs [48]. The next chapter discusses in more detail the methods that can be applied to determine, from respiratory data, the pharmacodynamic models for single drugs and combinations of drugs. IV.
Resting Measurements
The simplest and arguably the most clinically relevant [9] data measurement is the effect of a drug on resting ventilation and/or CO2 (either arterial or end-tidal). However, interpretation of this measurement becomes more complex due to many factors (in addition to the chemoreflexes) determining resting ventilation. The functioning of the hypercapnic chemoreflex feedback loop when CO2 is increased will tend to minimize the measured effect of the drug. The resting ventilation is determined by the intersection of the ventilatory response to CO2 with the metabolic hyperbola (Figures 4.1 and 4.2). The ventilatory response to CO2 is often characterized as linear above a threshold and thus has three, relatively independent factors: wakefulness drive (i.e., ventilation below the CO2 threshold), slope, and position (i.e., the CO2 intercept at zero ventilation) of the hypercapniainduced ventilation. Other parameterizations of the response have also been used [10,49]. Many factors (e.g., hypoxia, acid–base status, pain, arousal, etc. [13,50,51]) influence the specific value of each of these parameters and it is important that the control and drug conditions are identical except for the drug under study. When studying the respiratory effects of a drug, the changes in the resting values need to be supplemented with further information about specific drug effects on the elements of the controller. Most frequently these are measurements of the effects on the hypercapnic or hypoxic chemoreflexes. In clinical studies, it is difficult to control the many factors involved; particularly in patients, it is often difficult to make reliable measurements. In many clinical studies, then, the resting control of breathing, particularly
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Figure 4.2 Illustration of how an increase in the ‘wakefulness’ drive, caused by an arousal, can affect the resting ventilation. The control response has a resting ventilation at the intersection of the wakefulness drive and the chemoreflex response. The drug caused a decrease in the slope with a small right shift of the chemoreflex and a reduction (but not elimination) of the wakefulness drive such that the resting ventilation is now determined by the intersection of the metabolic hyperbola and the chemoreflex line. The arousal caused an increase in the wakefulness drive and an elimination of the small right shift. Note that even though there was no change in the reduction of the slope, the resting ventilation and PaCO2 returned almost back to normal.
in post-operative patients, is often summarized by simply measuring the oxygen saturation with pulse oximetry over time. These studies can be of great clinical interest since post-operative desaturation has been associated with myocardial ischemia and arrhythmias. However, these studies are greatly aided by, at the least, a qualitative measurement of ventilation [31] and sleep state [52] as done during polysomnography in the sleep laboratory. V.
Hypercapnic Ventilatory Response
The hypercapnic ventilatory response has been used for 100 years to assess the ventilatory response to drugs [53,54]. While originally only different levels of inspired CO2 were used, it soon became apparent that measurement or estimation of the arterial CO2 is also required. The development of the rebreathing technique by Read provided a relatively simple technique to measure the hypercapnic response quickly, and thus it rapidly
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became popular for many drug studies [22]. The response is usually parameterized with a slope and intercept obtained by linear regression of the breath-by-breath ventilation on the PETCO2 [55]. The steady-state technique (constant inspired CO2 concentration for a long enough period of time for the ventilation to reach steady-state) has evolved with control over the endtidal CO2 via manipulation of the inspired gas tension (dynamic end-tidal forcing—DEF) using automatic computer feedback control [56–58]. It is used to obtain step responses [57] or pseudo-rebreathing inputs [59] and leads to more sophisticated input function designs, thereby obtaining more accurate estimates of the chemoreflex loop parameters [60]. These DEF techniques have often been aimed at estimating both a peripheral and a central CO2 gain based on separation of the response times. While both techniques (steady-state and rebreathing) yield several characteristic parameters of the respiratory system that are related to underlying mathematical models, the primary parameter of interest for drug studies is often the overall CO2 gain—the increase in ventilation resulting from an increase in CO2. While Read’s original work indicated that the rebreathing and steady-state gains were identical [22], subsequent analysis [61] and work by others indicate that gain measured by rebreathing is larger than when measured by the steady-state technique [22,61–65] and more importantly, may also result in differences in the estimation of the effects of drugs on the hypercapnic response [66]. The essential features of the Read rebreathing test are that the subject begins the process with a vital capacity breath and then starts rebreathing from a small bag (4–6 l) filled with 7% CO2 in oxygen. The purpose of these maneuvers is to start the rebreathing at a value close to the mixed venous level and thus quickly obtain an equilibrium in mixed venous, lung, rebreathing bag, arterial, and brain tissue PCO2 [22,61]. Since external gas exchange no longer occurs, the rise in PETCO2 is independent of the level of ventilation, effectively opening the CO2 control feedback loop. The top panel in Figure 4.3 shows the continuous waveform from a single subject undergoing a Read rebreathing test. Note the vital capacity breath at the start of the rebreathing as well as the initial step in PETCO2 . There is a short equilibrium period and then the PETCO2 slowly increases at a rate determined by the subject’s metabolic rate and body CO2 buffering capacity. Thus, the CO2 stimulus can be characterized as an initial step of A (mm Hg) and a rate of rise of R (mm Hg min1) [59]. The differences in ventilation between the steady-state data and the rebreathing data at the same PETCO2 , and the resulting difference in the slope of the response, are illustrated in the bottom panel of Figure 4.3. The steady-state data (four periods of 8 min of elevated PETCO2 ) are shown by the open circles. The initial rebreathing data (filled squares) show an almost vertical increase in ventilation without a change in PETCO2 , representing the initial step and equilibrium period for the rebreathing. These points cannot be used in
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Figure 4.3 Continuous tidal volume and CO2 waveforms from a subject performing a rebreathing experiment (upper panel) and the regression fits to rebreathing data (squares) and steady-state data (open circles) (Data from Ref. 63).
the calculation of the response slope. The open squares then show the breath-to-breath rebreathing data. These points are used to calculate the rebreathing ventilatory response. Note that the rebreathing data show the expected right shift, but also that the slope is steeper. This increase in slope can be predicted from theoretical analysis [59,61,63,67] and is due to the increase in cerebral blood flow with hypercapnia reducing the arterialto-tissue CO2 gradient. If the gradient is reduced to zero and it remains
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constant throughout the rebreathing period, then the measured slope should approximate the slope of the tissue PCO2-to-ventilation relationship. However, it is difficult to verify that these conditions are, in fact, achieved and maintained [67]. This analysis also shows that the measured slope will depend on the size of the initial step, A, which can present difficulties when, in a pharmacological study, the starting PETCO2 is increased due to the ventilatory depression of the drug. Bourke and Warley directly compared the hypercapnic responses measured with the steady-state and rebreathing methods following two doses of morphine [66]. They found that the control slopes were higher when measured with the rebreathing method than with the steady-state method, and that after morphine the steady-state method showed a rightward shift and no change in slope while when measured with the rebreathing method, there was a decrease in the slope. Applying the theoretical analysis of Berkenbosch et al. [59], they determined that most of this change in slope was due to the smaller A after morphine due to the increase in PETCO2 . Their conclusion was that the specific opioid effect was a rightward shift of the hypercapnic ventilatory response without a change in sensitivity, and that this was best measured with a steady-state technique [66]. Similarly, Linton et al. found that while the slopes of the CO2 response determined by steadystate and rebreathing were identical under control conditions, acidosis and alkalosis had quite different effects on the response as measured by the two methods [64]. The rebreathing technique has been further refined by Duffin and colleagues [49,62,68,69], who have advocated preceding the rebreathing not with a single vital capacity breath, but rather with a period of hyperventilation that reduces the PETCO2 into the mid-20s mm Hg. Rebreathing is started and ventilation is maintained by the resting (wakefulness) ventilatory drive (see above) as the PETCO2 gradually rises. Once a threshold is reached, the ventilation then increases linearly with PETCO2 until a second breakpoint is reached, when the ventilation then increases faster due to a rapid increase in ventilatory frequency (Figure 4.4). This technique still provides an estimate of the hypercapnic response slope that is larger than the steady-state response [62]. While this method may provide further information about drug effects on different parts of the response (e.g., basal ventilation, threshold T1 and slope S 2), it has not been applied to many drug studies [70,71]. For most drug studies, the use of a steady-state technique is the most appropriate for determining central chemosensitivity. Since these tests are often performed in hyperoxia, the peripheral chemoreceptors contribute little to the overall ventilatory drive. Special test or data analysis needs to be applied if separation of the peripheral and central chemosensitivities is desired.
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Figure 4.4 Duffin’s modified rebreathing test in which the commencement of rebreathing is after a period of hyperventilation in which the PETCO2 is lowered to below 30 mm Hg (data segment not shown). Several parameters can be estimated from these experiments: The basal ventilation prior to the increase with hypercapnia; the PETCO2 threshold T1 at which ventilation first increases; the initial segment slope, S1; the second threshold when respiratory frequency starts to greatly increase, T 2; and the final augmented slope, S2 (Data from Ref. 49).
A single breath test of CO2 has been used as a clinical test to determine the sensitivity of the peripheral chemoreceptors [72]; this test has been applied in determining the effects of spinal anesthesia [73]. While this test is quick to perform, the signal-to-noise ratio is poor and the estimation of the peripheral gain can be contaminated with other
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factors [74]. Interestingly, the sensitivities of the single-breath CO2 test and the acute hypoxic response were not found to be correlated [75]. This test for peripheral sensitivity should not be used in place of an hypoxic test if the effect of a drug on the ventilatory response to hypoxia is desired. The main advantage of the rebreathing technique, aside from the relatively simple equipment required, is that a hypercapnic response can be obtained in several minutes, a fact which may be important if the drug effect is rapidly changing. The steady-state technique, if a constant inspired CO2 is used, may require over 10 min for ventilation to reach a steady state [64]. However, the technique of controlling the inspired CO2 such that the endtidal CO2 reaches a constant value immediately (e.g., step in PETCO2 [57]) or follows some other pre-described waveform (e.g., sinewave [76] or a timevarying binary sequence [60,77]) can permit a much shorter time period in which to make the ventilatory measurements. These shorter DEF steps can be combined with a two-compartment mathematical model of the ventilatory controller and the response separated into a fast (peripheral) and a slow (central) component [78,79]. However, from human experiments it cannot be certain that the dynamic component separation with statistical parameter estimation actually results in a central and peripheral component. Using an artificially perfused brainstem (ABP), spontaneously breathing cat preparation, DeGoede et al. directly compared the results of the DEF step response parameter estimation method with the determination of the central and peripheral gains from the ABP method [80]. Good agreement was found between the gains and the CO2 (extrapolated) threshold determined by these two methods over a range of peripheral gains. In carotid-body-resected human subjects, the DEF technique also found no peripheral component [81]. More recently, the DEF technique was used with a multifrequency binary sequence of PETCO2 in patients with unilateral and bilateral resections of the carotid bodies [82]. Interestingly, when using this technique, the bilaterally resected patients had lower sensitivity for the single slow component than the control subjects. Although the step input seems to satisfactorily separate the gains, its use has been criticized on the grounds that the input is not persistently exciting enough to obtain good estimates of the peripheral gain [60]. In essence, this is because the fast-component gain is only estimated during the initial rapid increase in ventilation, and the total gain is determined by the estimated final ventilation. A proposed solution to this problem is to use a waveform that consists of multiple up and down steps of different duration, resulting in a multi-frequency binary sequence (Figure 4.5, top panel). The number of transitions and the duration of the steps can be determined theoretically to provide an input that is appropriate for the assumed central and peripheral time constants [60]. A drawback of this technique is that an experimental period of over 20 min is required; however,
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Figure 4.5 Use of a specialized dynamic end-tidal CO2 forcing function to obtain the estimates of the effects of a drug on the parameters of the chemoreflex loops. The top panel shows a multifrequency binary sequence (theoretical) in end-tidal CO2. This input function is used to excite the ventilatory controller (bottom panel) before and after propofol administration. This waveform allows for the parameters of both fast (peripheral) and slow (central) chemoreflex loops to be estimated from the ventilatory response. This divides the response into central, Vc, and peripheral, Vp, components (Data from Ref. 83).
it provides a more reliable and accurate estimate of the peripheral gain. This technique has been applied to drug studies in which the drug level was constant for the time period, allowing estimation of changes in the central and peripheral CO2 gains (Figure 4.5, bottom panel) [83]. Besides understanding the effects of a drug on the hypercapnic response when the drug concentration is constant, it is also of interest to study the time course of the drug effect. For many drugs that have slowly changing respiratory effects, repeated hypercapnic studies can be made, but with drugs that produce rapid alterations in ventilation (e.g., anesthetic induction drugs), other techniques must be employed. An obvious method
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is to hold the end-tidal CO2 constant by adjusting the inspired CO2 as ventilation changes with the drug effect. Since respiratory depressants are usually studied, it is necessary to start at a hypercapnic level so that the inspired CO2 can be lowered as the ventilation declines. By repeating the experiment twice, the so-called dual iso-hypercapnic technique, the time course of the change in the hypercapnic sensitivity can be constructed [84]. The major drawback to this technique is the need to give two drug doses, most likely on different days, unless the drug has very fast pharmacokinetics (e.g., remifentanil [85]). The day-to-day variability in the hypercapnic response will greatly diminish the ability to accurately measure changes in the hypercapnic sensitivity. Bouillon and coworkers have applied an indirect response model to the ventilatory depression caused by alfentanil [10] and remifentanil [86] in the non-steady state. This modeling technique permits the prediction of the time course of respiratory depression after a bolus dose of the drug, considering the indirect effect of the slow rise in CO2-stimulating ventilation [87]. While this technique is attractive and provides a clinically relevant model of druginduced respiratory depression, it requires estimation of the hypercapnic response for each subject prior to the drug experiment. The method also requires assumptions on the shape of the hypercapnic response, particularly in the transient, when CO2 is less than its final steady-state value, e.g., below the metabolic hyperbola (Figure 4.2). This model has not been fully experimentally validated and has the additional drawback of not being able to predict the apnea that is observed clinically at high drug doses [88]. VI.
Hypoxic Ventilatory Response
Although the ventilatory response to hypoxia has been described since the early part of the 20th century, the effect of morphine on the hypoxic response was not studied until 1975 [89]. Although hypoxia is a common and dangerous clinical condition, the effects of drugs on the hypoxic response have been difficult to study. The hypoxic response is a nonlinear function of the arterial O2 [90], is strongly affected by the CO2 level [91,92], and has a time-dependent effect [93]. The time dependence of the ventilatory response can be seen as a pronounced decrease in ventilation (hypoxic ventilatory decline—HVD) after 5–15 min of isocapnic hypoxia [90,94,95]. In studying the hypoxic response, the negative feedback from the increased ventilation lowering the CO2 and raising the O2 has to be eliminated. Thus isocapnic conditions have to be maintained not only throughout the time period of the hypoxic stimulus, but also during the different drug conditions. It is important that the CO2 chosen be high enough to ensure that the hypercapnic chemoreflex is active. Since many of the tested drugs will cause hypercapnia at rest, the control hypoxic
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responses need to be made at similar hypercapnic levels. While a rebreathing type of hypoxic test has been advocated [96], the biphasic nature of the hypoxic response can readily affect ventilation before the end of the rebreathing period. Thus, hypoxic responses should be tested with a DEF-type system so that PETCO2 can be controlled on a breath-to-breath basis and precise control over PETO2 (and saturation) can be maintained. However, since in the clinical situations PETCO2 will obviously not be controlled, poikilocapnic hypoxic tests have been advocated as a better measurement of the clinical effects of drugs [97–99]. The results for a drug effect may be different depending on whether an isocapnic or a poikilocapnic hypoxic test is used. For example, Sjogen et al. found that the isocapnic acute hypoxic response was 4.7 2.3 l min1 but only 1.4 1.0 l min1 for the poikilocapnic response; 0.6 MAC of isoflurane did not significantly reduce the poikilocapnic response (1.3 0.8 l min1) but did reduce the isocapnic response (2.3 1.4 l min1, p 5 0.02) [98]. The first issue in quantifying the hypoxic response is to determine the parameters of the nonlinear relationship between ventilation and arterial PO2 (Figure 4.6). There is no underlying physiological basis for any particular parameterization and the cellular mechanisms underlying the non-linear response are poorly understood (see Nurse, Chapter 1, this volume). Either a hyperbola [100–102] or linear-in-saturation [96] model has been the most commonly used. The availability of pulse oximetry has made the use of the linear-in-saturation model popular. Several studies have compared the fit of the various models to the initial hypoxic response (note that it is not appropriate to characterize this as the steady-state hypoxic response since the development of the acute response and HVD overlap, and no real steady state is seen in the first 20–30 min of hypoxia [93]). The studies have not found any significant difference in the fit between the hyperbolic equation and the linear-insaturation equations [103,104]. Figure 4.6 shows a comparison of seven models to the hypoxic response of a single subject. By visual inspection, it is readily apparent that the variability in the data makes all the models, except for a direct linear fit to PETCO2 , seem to be suitable. Because the linear slope with respect to saturation is the most readily interpretable, it is the most common model used to quantify a drug effect (expressed in units of l min1 % sat1, while noting that since the ventilation increases with decreasing saturation, this gain is negative, but is commonly expressed as the change in ventilation for a decrease in saturation giving a positive value). There are no studies of whether the shape of the acute hypoxic response is altered by a drug, but given the variability of the data it is doubtful that minor changes could be detected.
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Figure 4.6 Examples of different model fits to the hypoxic ventilatory response in one subject. Top and center panels show the fit to PETO2 and the bottom panel shows the fit to the measured saturation (Data from Ref. 103).
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The ventilatory response to sustained hypoxia has several time domains, and therefore it is not possible to define a true steady-state response [93]. The response over the first 20 min of hypoxia is markedly biphasic, with ventilation declining by approximately 50% of the initial rise over this period of time [95]. This response has been modeled in a way that is similar to the dynamic hypercapnic response model, with two exponential terms but with one stimulatory and the other inhibitory [105–107]. However, few drug studies have examined specific changes in these model parameters, but rather used pseudo-steady-state methods of parameterizing the hypoxic response. These methods make the assumption that the ventilation at 3–5 min after the onset of hypoxia is not affected by HVD and that an average ventilation during this time period will be a measure of the peripheral (carotid body) chemoreflex. The ventilation occurring between 15 and 20 min of hypoxia will reflect the development of HVD and can be used to calculate the sustained ventilatory response. An interesting characteristic of HVD is that a brief period of normoxia (e.g., 5 min) will not reverse the depression [108]. The magnitude of this repeated hypoxic response, when compared with the initial hypoxic response, can be used as a measure of HVD. A drug’s effect can be assessed both on the acute hypoxic response and HVD [109]. Figure 4.7 shows how the characteristics of the hypoxic response can be calculated. A drug may have independent effects on the acute response or HVD. However, it is important to understand how HVD is assessed. The amount of HVD is not fixed, but rather depends on the magnitude of the acute response [92,110]. Thus, a drug that reduced the acute response would also seem to increase the amount of HVD if measured only by the sustained hypoxic sensitivity. In drug effect studies, it is better to express the magnitude of HVD by the ratio of the increase in ventilation from the baseline to the acute peak to the amount of ventilatory decline from the acute peak ventilation to the ventilation after 15–20 min of hypoxia. These changes in ventilation can be normalized by the saturation at these time points if the saturation is not held adequately constant throughout the whole hypoxic period. While ventilation is most commonly used as the measured variable, it is important to note that HVD seems to occur primarily in tidal volume, and separating the response into tidal volume and ventilatory frequency may reveal an effect on one or the other [111]. VII.
Changes in Airway Pressure
Since airway obstruction is a common occurrence after many respirationdepressing drugs [112], there has been development of techniques to
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Figure 4.7 Response to 20 min of isocapnic hypoxic and a repeat hypoxia exposure after 5 min of normoxia before (top panel) and after alfentanil (bottom panel). The acute hypoxic response for the first step (intervals 1 and 2) and the second step (intervals 4 and 5) can be calculated. The HVD can be calculated as the ratio of these acute steps or as the absolute difference between the ventilation during intervals 2 and 3. Normalizing the change in ventilation by the change in saturation results in calculation of hypoxic sensitivities (Data from Ref. 109).
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quantify this effect. This can be done in several ways, either through an added resistance or through the application of negative airway pressure. The former studies have been developed to simulate the effects of airway disease while the latter, to simulate obstructive sleep apnea (see Chapter 10 by Schwartz, this volume). In an awake subject, the application of negative airway pressure activates compensating activity in the pharyngeal dilator muscles; complete airway obstruction seldom occurs with negative pressures even as low as 30 cm H2O. However, during sleep, a negative nasal pressure of 10 cm H2O can induce a syndrome-like obstructive sleep apnea in normal subjects [113], and a positive pressure is required to maintain a patent airway in patients with obstructive sleep apnea. Application of different airway pressures (from positive to negative) has been used to test for changes caused by diazepam [114], midazolam [115], isoflurane [116] and vecuronium [117]. Typically, the outcome measurement in these studies is the pressure that causes airway collapse (critical pressure, Pcrit), measured either directly [115] or by extrapolation from the flow versus airway pressure on flow-limited breaths [116]. The measurement of flow limitation requires the measurement of esophageal pressure; the additional measurement of genioglossial EMG may also be useful (Figure 4.8). While a lower Pcrit in anesthetized patients correlated with the severity of sleep-disordered breathing [118], it is unknown how the drug-induced propensity for airway obstruction as measured by Pcrit correlates with clinically important outcomes. VIII.
Other Stimuli
Pain and exercise are other common stimuli that have pronounced ventilatory effects. The ventilatory response to exercise has been extensively studied throughout the 20th century without complete understanding of all the mechanisms involved. This makes it difficult to interpret any drug effects on the exercised induced ventilatory responses. Two studies have found that opioids do little to blunt the link between exercise and ventilation even at doses that caused considerable resting hypercapnia [119,120]. Since painful stimuli can only be quantified subjectively and introducing pain is an additional variable in drug effect studies, the interpretation of the results of these studies can be quite complex. Surgical skin incision has been studied for its effects on ventilation and ventilatory pattern [121,122]. The interaction of painful stimuli and drug-induced depression of the chemoreflexes has also been studied for both the hypercapnic and hypoxic responses. Pain seems to have little effect on the control response [123] and does not counter the effects of inhalational anesthetics on the hypoxic response [124]. When studies involve the use of painful stimuli it is important to consider anxiety and arousal induced by
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Flow Pm
expiratory port
EMGgg
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2
B
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E
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-20
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MTA EMGgg 20 sec
Figure 4.8 Top panel show experimental setup to change the airway pressure (both positive and negative) to test the effects of isoflurane on the collapsibility of the upper airway. Bottom panel shows the physiological response during isoflurane anesthesia when the airway pressure is reduced from þ20 cm H2O (A) to levels that cause flow limitation (B) or complete obstruction (C). Note that even a brief pulse of 30 cm H2O (D) did not activate any EMGgg even though there is a robust signal when the subject is awake (E) and is asked to protrude the tongue (F). Pm—mask pressure; Pnp—nasopharyngeal pressure; Pop—oropharyngeal pressure; Php—hypopharyngeal pressure; Pes—esophageal pressure; Piso—isoflurane concentration; EMGgg—genioglossus electromyogram; MTA—moving time average (Data from Ref. 116).
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the stimuli as well as the specific type of stimuli employed (e.g., electrical [123], pressure [125] or thermal [126]). References 1.
2.
3.
4.
5. 6.
7.
8. 9. 10.
11. 12. 13. 14. 15.
Bailey, P.L., Streisand, J.B., East, K.A., East, T.D., Isern, S., Hansen, T.W., Posthuma, E.F.M., Tozendaal, F.W., Pace, N.L. and Stanley, T.H., Differences in magnitude and duration of opioid-induced respiratory depression and analgesics with fentanyl and sufentanil, Anesth. Analg. 70, 8–15, 1990. Kagawa, S., Stafford, M.J., Waggener, T.B. and Severinghaus, J.W., No effect of naloxone on hypoxia-induced ventilatory depression in adults, J. Appl. Physiol. 52: 1030–1034, 1982. Cummin, A.R.C., Jacobi, M.S., Patil, C.P., Telford, R.J., Morgan, C.N. and Saunders, K.B., The effect of almitrine on the steady-state ventilatory response to carbon dioxide at rest and during exercise in man, Eur. Respir. J. 3, 693–698, 1990. Daskalopoulou, E., Patakas, D., Tsara, V., Zoglopitis, F. and Maniki, E., Comparison of almitrine bismesylate and medroxyprogesterone acetate on oxygenation during wakefulness and sleep in patients with chronic obstructive lung disease, Thorax 45, 666–669, 1990. Calverley, P.M.A., Robson, R.H. and Wraith, P.K., The ventilatory effects of doxapram in normal man, Clin. Sci. 65, 65–69, 1983. Amin, H.M., Sopchak, A.M., Foss, J.F., Esposito, B.F., Roizen, M.F. and Camporesi, E.M., Efficacy of methylnaltrexone versus naloxone for reversal of morphine-induced depression of hypoxic ventilatory response, Anesth. Analg. 78, 701–705, 1994. Flo¨gel, C.M., Ward, D.S., Wada, D.R. and Ritter, J.W., The effects of largedose flumazenil on midazolam-induced ventilatory depression, Anesth. Analg. 77, 1207–1214, 1993. Borison, H.L., Central nervous respiratory depressants—control-systems approach to respiratory depression, Pharmacol. Ther. [B] 3, 211–226, 1977. Knill, R.L., Wresting or resting ventilation, Anesthesiology 59, 599–600, 1983. Bouillon, T., Schmidt, C., Garstka, G., Heimbach, D., Stafforst, D., Schwilden, H. and Hoeft, A., Pharmacokinetic–pharmacodynamic modeling of the respiratory depressant effect of alfentanil, Anesthesiology 91, 144–155, 1999. Dejours, P., Chemoreflexes in breathing, Physiol. Rev. 42, 335–358, 1962. Fink, B.R., Influence of cerebral activity in wakefulness on regulation of breathing, J. Appl. Physiol. 16, 15–20, 1961. Ward, D.S. and Karan, S., Effects of pain and arousal on the control of breathing, J. Anesth. 16, 216–221, 2002. Orem, J. and Trotte, R.H., Behavioral control of breathing, News Physiol. Sci. 9, 228–232, 1994. Guz, A., Brain, breathing and breathlessness, Respir. Physiol. 109, 197–204, 1997.
Measurement of Drug Effects on Ventilatory Control 16. 17.
18.
19.
20.
21.
22. 23. 24.
25.
26.
27.
28.
29.
30. 31.
125
Berthon-Jones, M. and Sullivan, C.E., Ventilatory and arousal responses to hypoxia in sleeping humans, Am. Rev. Respir. Dis. 125, 632–639, 1982. Corfield, D.R., Roberts, C.A., Griffiths, M.J. and Adams, L., Sleep-related changes in the human ‘neuromuscular’ ventilatory response to hypoxia, Respir. Physiol. 117, 109–120, 1999. Knill, R.L., Moote, C.A., Skinner, M.I., Rose, E.A. and Lok, P.Y.K., Morphine-induced ventilatory depression is potentiated by non-REM sleep, Can. J. Anaesth. 34, S101–S102, 1987. Forrest, W.H., Jr. and Bellville, J.W., The effect of sleep plus morphine on the respiratory response to carbon dioxide, Anesthesiology 25, 137–141, 1964. Grodins, F., Buell, J. and Bart, A.J., Mathematical analysis and digital simulation of the respiratory control system, J. Appl. Physiol. 22, 260–276, 1967. Khoo, M.C.K. and Yamashiro, S., Models of the control of breathing, in Respiratory Physiology. An Analytic Approach, Chang, H.K. and Paiva, M., eds., New York, Marcel Dekker, Inc., pp. 799–829, 1989. Read, D.J.C., A clinical method for assessing the ventilatory response to carbon dioxide, Australas. Ann. Med. 16, 20–32, 1967. Friedman, L.M., Furberg, C.D. and deMets, D.L., Fundamentals of Clinical Trials, 3rd edn., New York, Springer-Verlag, 1998. Jenkins, J.S., Valcke, C.P. and Ward, D.S., A programmable system for acquisition and reduction of respiratory physiology data, Ann. Biomed. Eng. 17, 93–108, 1989. Severinghaus, J.W., Water vapor calibration errors in some capnometers: Respiratory conventions misunderstood by manufacturers? Anesthesiology 70, 996–998, 1989. Milic-Emili, J., Grassino, A.E. and Whitelaw, W.A., Measurement and testing of respiratory drive, in Regulation of Breathing, Hornbein, T.F., ed., New York, Marcel Dekker, Inc., pp. 675–743, 1981. Rebuck, A.S. and Slutsky, A.S., Measurement of ventilatory responses to hypercapnia and hypoxia, in Regulation of Breathing, Hornbein, T.F., ed., Vol. 17, Part 2, New York, Marcel Dekker, Inc., pp. 745–772, 1981. Akiyama, Y. and Kawakami, Y., Clinical assessment of the respiratory control system, in Control of Breathing in Health and Disease, Altose, M.D. and Kawakami, Y., eds., New York, Marcel Dekker, Inc., pp. 251–287, 1999. Cherniack, N.S., Dempsey, J., Fencl, V., Fitzgerald, R.S., Lourenco, R.V., Rebuck, A.S., Rigg, J., Severinghaus, J.W., Weil, J.W., Whitelaw, W.A. and Zwilich, C.W., Workshop on assessment of respiratory control in humans. I. Methods of measurement of ventilatory responses to hypoxia and hypercapnia, Am. Rev. Respir. Dis. 115, 177–181, 1977. Severinghaus, J.W. and Naifeh, K.H., Accuracy of response of six pulse oximeters to profound hypoxia, Anesthesiology 67, 551–558, 1987. Rosenberg, J., Rasmussen, G.I., Wojdemann, K.R., Kirkeby, L.T., Jorgensen, L.N. and Kehlet, H., Ventilatory pattern and associated episodic hypoxaemia in the late postoperative period in the general surgical ward, Anaesthesia 54, 323–328, 1999.
126 32.
33.
34.
35.
36.
37. 38. 39.
40.
41.
42.
43.
44.
45.
Ward Mo¨ller, J.T., Johannessen, N.W., Espersen, K., Ravlo, O., Pedersen, B.D., Jensen, P.F., Rasmussen, N.H., Rasmussen, L.S., Pedersen, T. and Cooper, J.B., et al., Randomized evaluation of pulse oximetry in 20,802 patients: II. Perioperative events and postoperative complications, Anesthesiology 78, 445–453, 1993. Reeder, M.K., Goldman, M.D., Loh, L., Muir, A.D., Casey, K.R. and Lehane, J.R., Late postoperative nocturnal dips in oxygen saturation in patients undergoing major abdominal vascular surgery. Predictive value of preoperative overnight pulse oximetry, Anaesthesia 47, 110–115, 1992. Wheatley, R.G., Somerville, I.D., Sapsford, D.J. and Jones, J.G., Postopertive hypoxaemia: Comparison of extradural, I.M. and patient-controlled opioid analgesia, Br. J. Anaesth. 64, 267–275, 1990. Catley, D.M., Thorton, M., Jordan, C., Tech, B., Lehane, J.R., Royston, D. and Jones, J.G., Pronounced, episodic oxygen desaturation in the postoperative period: Its association with ventilatory pattern and analgesic regimen, Anesthesiology 63, 20–28, 1985. Moller, J.T., Cluitmans, P., Rasmussen, L.S., Houx, P., Rasmussen, H., Canet, J., Rabbitt, P., Jolles, J., Larsen, K. and Hanning, C.D., Long-term postoperative cognitive dysfunction in the elderly: ISPOCD1 study, Lancet 351, 857–861, 1998. Prabhakar, N.R., Sleep apneas. An oxidative stress? Am. J. Respir. Crit. Care Med. 165, 859–860, 2002. Leung, R.S.T. and Bradley, T.D., Sleep apnea and cardiovascular disease, Am. J. Respir. Crit. Care Med. 164, 2147–2165, 2001. Konno, K. and Mead, J., Measurement of the separate volume changes of rib cage and abdomen during breathing, J. Appl. Physiol. 22, 407–422, 1967. Perez, W. and Tobin, M.J., Separation of factors responsible for change in breathing pattern induced by instrumentation, J. Appl. Physiol. 59, 1515–1520, 1985. Gilbert, R., Auchincloss, J.H., Jr., Brodsky, J. and Boden, W., Changes in tidal volume, frequency, and ventilation induced by their measurement, J. Appl. Physiol. 33, 252–254, 1972. Litman, R.S., Kottra, J.A., Gallagher, P.R. and Ward, D.S., Diagnosis of anesthetic-induced upper airway obstruction in children using respiratory inductance plethysmography, J. Clin. Monit. Comput. 17, 279–285, 2002. Brown, K., Aun, C., Jackson, E., Mackersie, A., Hatch, D. and Stocks, J., Validation of respiratory inductive plethysmography using the Qualitative Diagnostic Calibration method in anaesthetized infants, Eur. Respir. J. 12, 935–943, 1998. Brown, K., Aun, C., Stocks, J., Jackson, E., Mackersie, A. and Hatch, D., A comparison of the respiratory effects of sevoflurane and halothane in infants and young children, Anesthesiology 89, 86–92, 1998. Warner, D.O., Warner. M.A. and Ritman, E.L., Human chest wall function while awake and during halothane anesthesia. I. Quiet breathing, Anesthesiology 82, 6–19, 1995.
Measurement of Drug Effects on Ventilatory Control 46.
47.
48.
49.
50.
51. 52.
53. 54. 55. 56.
57. 58. 59.
60.
61. 62.
63.
127
Benameur, M., Goldman, M.D., Ecoffey, C. and Gaultier, C., Ventilation and thoracoabdominal asynchrony during halothane anesthesia in infants, J. Appl. Physiol. 74, 1591–1596, 1993. Mildh, L.H., Scheinin, H. and Kirvela, O.A., The concentration-effect relationship of the respiratory depressant effects of alfentanil and fentanyl, Anesth. Analg. 93, 939–946, 2001. Bragg, P., Zwass, M.S., Lau, M. and Fisher, D.M., Opioid pharmacodynamics in neonatal dogs: Differences between morphine and fentanyl, J. Appl. Physiol. 79, 1519–1524, 1995. Duffin, J., Mohan, R.M., Vasiliou, P., Stephenson, R. and Mahamed, S., A model of the chemoreflex control of breathing in humans: Model parameters measurement, Respir. Physiol. 120, 13–26, 2000. Shea, S.A., Walter, J., Pelley, C., Murphy, K. and Guz, A., The effect of visual and auditory stimuli upon resting ventilation in man, Respir. Physiol. 68, 345–357, 1987. Shea, S.A., Behavioural and arousal-related influences on breathing in humans, Exp. Physiol. 81, 1–26, 1996. Rosenberg, J., Wildschiodtz, G., Pedersen, M.H,, von Jessen, F. and Kehlet, H., Late postoperative nocturnal episodic hypoxaemia and associated sleep pattern, Br. J. Anaesth. 72, 145–150, 1994. Loewy, A., Zur kenntniss der erregbarkeit des athemcentrums, Archiv fur die Gesammte Physiologie 47, 601–621, 1890. Lindhard, J., On the excitibility of the respiratory centre, J. Physiol. (Lond.) 42, 337–358, 1911. Goodman, N.W. and Curnow, J.S.H., The ventilatory response to carbon dioxide, Br. J. Anaesth. 57, 311–318, 1985. Robbins, P.A., Swanson, G.D. and Howson, M.G., A prediction–correction scheme for forcing alveolar gases along certain time courses, J. Appl. Physiol. 52, 1353–1357, 1991. Swanson, G.D. and Bellville, J.W., Step changes in end-tidal CO2: Methods and implications, J. Appl. Physiol. 39, 377–385, 1975. Kawakami, Y., Yoshikawa, T., Asanuma, Y. and Murao, M., A control system for arterial blood gases, J. Appl. Physiol. 50, 1362–1366, 1981. Berkenbosch, A., DeGoede, J., Olievier, C.N. and Schuitmaker, J.J., A pseudo-rebreathing technique for assessing the ventilatory response to carbon dioxide in cats, J. Physiol. (Lond.) 381, 483–495, 1986. Pedersen, M.E.F., Fatemian, M. and Robbins, P.A., Identification of fast and slow ventilatory responses to carbon dioxide under hypoxic and hyperoxic conditions in humans, J. Physiol. (Lond.) 521, 273–287, 1999. Read, D.J.C. and Leigh, J., Blood-brain tissue PCO2 relationships and ventilation during rebreathing, J. Appl. Physiol. 23, 53–70, 1967. Mohan, R.M., Amara, C.E., Cunningham, D.A. and Duffin, J., Measuring central-chemoreflex sensitivity in man: Rebreathing and steady-state methods compared, Respir. Physiol. 115, 23–33, 1999. Berkenbosch, A., Bovill, J.G., Dahan, A., DeGoede, J. and Olievier, I.C.W., The ventilatory CO2 sensitivities from Read’s rebreathing method and the
128
64.
65.
66.
67.
68. 69.
70.
71.
72.
73.
74. 75.
76. 77. 78.
79.
Ward steady-state method are not equal in man, J. Physiol. (Lond.) 411, 367–377, 1989. Linton, R.A,, Poole-Wilson, P.A., Davies, R.J. and Cameron, I.R., A comparison of the ventilatory response to carbon dioxide by steady-state and rebreathing methods during metabolic acidosis and alkalosis, Clin. Sci. Mol. Med. Suppl. 42, 239–249, 1973. Jacobi, M.S., Patil, C.P. and Saunders, K.B., Transient, steady-state and rebreathing responses to carbon dioxide in man, at rest and during light exercise, J. Physiol. (Lond.) 411, 85–96, 1989. Bourke, D.L. and Warley, A., The steady-state and rebreathing methods compared during morphine administration in humans, J. Physiol. (Lond.) 419, 509–517, 1989. Dahan, A., Berkenbosch, A., DeGoede, J., Olievier, I.C.W. and Bovill, J.G., On a pseudo-rebreathing technique to assess the ventilatory sensitivity to carbon dioxide in man, J. Physiol. (Lond.) 423, 615–629, 1990. Duffin, J. and McAvoy, G.V., The peripheral-chemoreceptor threshold to carbon dioxide in man, J. Physiol. (Lond.) 406, 15–26, 1988. Mohan, R.M., Amara, C.E., Vasiliou, P., Corriveau, E.P., Cunningham, D.A. and Duffin, J., Chemoreflex model parameters measurement, Adv. Exp. Med. Biol. 450, 185–193, 1998. Katzman, M.A., Duffin, J., Shlik, J. and Bradwejn, J., The ventilatory response to cholecystokinin tetrapeptide in healthy volunteers, Neuropsychopharmacology 26, 824–831, 2002. Vovk, A., Duffin, J., Kowalchuk, J.M., Paterson, D.H. and Cunningham, D.A., Changes in chemoreflex characteristics following acute carbonic anhydrase inhibition in humans at rest, Exp. Physiol. 85, 847–856, 2000. McClean, P.A., Phillipson, E.A., Martinez, D. and Zamel, N., Single breath of CO2 as a clinical test of the peripheral chemoreflex, J. Appl. Physiol. 64, 84–89, 1988. Steinbrook, R.A. and Concepcion, M., Respiratory effects of spinal anesthesia: Resting ventilation and single-breath CO2 response, Anesth. Analg. 72, 182–186, 1991. Khoo, M.C.K., A model-based evaluation of the single-breath CO2 ventilatory response test, J. Appl. Physiol. 68, 393–399, 1990. Chua, T.P. and Coats, A.J., The reproducibility and comparability of tests of the peripheral chemoreflex: Comparing the transient hypoxic ventilatory drive test and the single-breath carbon dioxide response test in healthy subjects, Eur. J. Clin. Investig. 25, 887–892, 1995. Daubenspeck, J.A., Frequency analysis of CO2 regulation: Afferent influences on tidal volume control, J. Appl. Physiol. 35, 662–672, 1973. Yang, F. and Khoo, M.C., Ventilatory response to randomly modulated hypercapnia and hypoxia in humans, J. Appl. Physiol. 76, 2216–2223, 1994. Ward, D.S. and Bellville, J.W., Effect of intravenous dopamine on hypercapnic ventilatory response in human, J. Appl. Physiol. 55, 1418–1425, 1983. Dahan, A., van den Elsen, M.J.L.J., Berkenbosch, A., DeGoede, J., Olievier, I.C.W., van Kleef, J. and Bovill, J.G., Effects of subanesthetic halothane on the
Measurement of Drug Effects on Ventilatory Control
80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
90.
91. 92. 93. 94. 95.
129
ventilatory responses to hypercapnia and acute hypoxia in healthy volunteers, Anesthesiology 80, 727–738, 1994. DeGoede, J., Berkenbosch, A., Ward, D.S., Bellville, J.W. and Olievier, C.N., Comparison of chemoreflex gains obtained with two different methods in cats, J. Appl. Physiol. 59, 170–179, 1985. Bellville, J.W., Whipp, B.J., Kaufman, R.D., Swanson, G.D., Aqleh, K.A. and Wiberg, D.M., Central and peripheral chemoreflex loop gain in normal and carotid body-resected subjects, J. Appl. Physiol. 46, 843–853, 1979. Fatemian, M., Nieuwenhuijs, D.J.F., Teppema, L.J., Meinesz, S., van der Mey, A.G.L., Dahan, A. and Robbins, P.A., The respiratory response to carbon dioxide in humans with unilateral and bilateral resections of the carotid bodies, J. Physiol. (Lond.) 549, 965–973, 2003. Nieuwenhuijs, D., Sarton, E., Teppema, L.J., Kruyt, E., Olievier, I., van Kleef, J. and Dahan, A., Respiratory sites of action of propofol: Absence of depression of peripheral chemoreflex loop by low-dose propofol, Anesthesiology 95, 889–895, 2001. Blouin, R.T., Conrad, P.F. and Gross, J.B., Time course of ventilatory depression following induction doses of propofol and thiopentathal, Anesthesiology 75, 940–944, 1991. Babenco, H.D., Conard, P.F. and Gross, J.B., The pharmacodynamic effect of a remifentanil bolus on ventilatory control, Anesthesiology 92, 393–398, 2000. Bouillon, T., Bruhn, J., Radu-Radulescu, L., Andresen, C., Cohane, C. and Shafer, S.L., A model of the ventilatory depressant potency of remifentanil in the non-steady state, Anesthesiology 99, 779–787, 2003. Jusko, W.J. and Ko, H.C., Physiologic indirect response models characterize diverse types of pharmacodynamic effects, Clin. Pharmacol. Ther. 56, 406–419, 1994. Gross, J.B., When you breathe IN you inspire, when you DON’T breathe, you . . . expire: New insights regarding opioid-induced ventilatory depression, Anesthesiology 99, 767–770, 2003. Weil, J.V., McCullough, R.E., Kline, J.S. and Sodal, I.E., Diminished ventilatory response to hypoxia and hypercapnia after morphine in normal man, N. Engl. J. Med. 292, 1103–1106, 1975. Bisgard, G.E. and Neubauer, J.A., Peripheral and central effects of hypoxia, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, Inc., pp. 617–668, 1995. Rebuck, A.S. and Woodley, W.E., Ventilatory effects of hypoxia and their dependence on PCO2, J. Appl. Physiol. 38, 16–19, 1975. Easton, P.A. and Anthonisen, N.R., Carbon dioxide effects on the ventilatory response to sustained hypoxia, J. Appl. Physiol. 64, 1451–1456, 1988. Powell, F.L., Milsom, W.K. and Mitchell, G.S., Time domains of the hypoxic ventilatory response, Respir. Physiol. 112, 123–134, 1998. Weiskopf, R.B. and Gabel, R.A., Depression of ventilation during hypoxia in man, J. Appl. Physiol. 39, 911–915, 1975. Easton, P.A., Slykerman, L.J. and Anthonisen, N.R., Ventilatory response to sustained hypoxia in normal adults, J. Appl. Physiol. 61, 906–911, 1986.
130
Ward
96.
Rebuck, A.S. and Campbell, J.M., A clinical method for assessing the ventilatory response to hypoxia, Am. Rev. Respir. Dis. 109, 345–350, 1974. Sjogren, D., Sollevi, A., Ebberyd, A. and Lindahl, S.G.E., Poikilocapnic hypoxic ventilatory response in humans during 0.85 MAC isoflurane anesthesia, Acta Anaesthesiol. Scand. 38, 149–155, 1994. Sjogren, D., Sollevi, A., Ebberyd, A. and Lindahl, S.G., Isoflurane anaesthesia (0.6 MAC), hypoxic ventilatory responses in humans, Acta. Anaesthesiol. Scand. 39, 17–22, 1995. Sjogren, D., Lindahl, S.G., Gottlieb, C. and Sollevi, A., Ventilatory responses to acute and sustained hypoxia during sevoflurane anesthesia in women, Anesth. Analg. 89, 209–214, 1999. Lloyd, B.B., Jukes, M.G.M. and Cunningham, D.J.C., The relation between alveolar oxygen pressure and the respiratory response to carbon dioxide in man, Q. J. Exp. Physiol. 43, 214–227, 1958. Weil, J.V., Bryne-Quinn, E., Sodal, I.E., Friesen, W.O., Underhill, B., Filley, G.F. and Grover, R.F., Hypoxic ventilatory drive in normal man, J. Clin. Investig. 49, 1061–1072, 1970. Weil, J.V. and Zwillich, C.W., Assessment of ventilatory response to hypoxia: Methods and interpretation, Chest 70, 124–128, 1976. van Klaveren, R.J. and Demedts, M., A mathematical and physiological evaluation of the different hypoxic response models in normal man, Respir. Physiol. 113, 123–133, 1998. Kronenberg, R., Hamilton, F.N., Gabel, R., Hickey, R., Read, D.J.C. and Severinghaus, J.W., Comparison of three methods for quantitating respiratory response to hypoxia in man, Respir. Physiol. 16, 109–125, 1972. Smith, W.D., Poulin, M.J., Paterson, D.H. and Cunningham, D.A., Dynamic ventilatory response to acute isocapnic hypoxia in septuagenarians, Exp. Physiol. 86, 117–126, 2001. Ward, D.S., Dahan, A. and Mann, C.B., Modelling the dynamic ventilatory response to hypoxia in humans, Ann. Biomed. Eng. 20, 181–194, 1992. Painter, R., Khamnei, S. and Robbins, P., A mathematical model of the human ventilatory response to isocapnic hypoxia, J. Appl. Physiol. 74, 2007–2015, 1993. Easton, P.A., Slykerman, L.J. and Anthonisen, N.R., Recovery of the ventilatory response to hypoxia in normal adults, J. Appl. Physiol. 64, 521–528, 1988. Cartwright, C.R., Henson, L.C. and Ward, D.S., Effects of alfentanil on the ventilatory response to sustained hypoxia, Anesthesiology 89, 612–619, 1998. Dahan, A., Ward, D.S, van den Elsen, M., Temp, J. and Berkenbosch, A., Influence of reduced carotid body drive during sustained hypoxia on hypoxic depression of ventilation in humans, J. Appl. Physiol. 81, 565–572, 1996. Easton, P.A. and Anthonisen, N.R., Ventilatory response to sustained hypoxia after pretreatment with aminophylline, J. Appl. Physiol. 64, 1445–1450, 1988. Hillman, D.R., Platt, P.R. and Eastwood, P.R., The upper airway during anaesthesia, Br. J. Anaesth. 91, 31–39, 2003.
97.
98.
99.
100.
101.
102. 103.
104.
105.
106. 107.
108.
109.
110.
111.
112.
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113. King, E.D., O’Donnell, C.P., Smith, P.L. and Schwartz, A.R., A model of obstructive sleep apnea in normal humans. Role of the upper airway, Am. J. Respir. Crit. Care Med. 161, 1979–1984, 2000. 114. Philip-Joet, F., Marc, I. and Series, F., Effects of genioglossal response to negative airway pressure on upper airway collapsibility during sleep, J. Appl. Physiol. 80, 1466–1474, 1996. 115. Litman, R.S., Hayes, J.L., Basco, M.G., Schwartz, A.R., Bailey, P.L. and Ward, D.S., Use of dynamic negative airway pressure (DNAP) to assess sedative-induced upper airway obstruction, Anesthesiology 96, 342–345, 2002. 116. Eastwood, P.R., Szollosi, I., Platt, P.R. and Hillman, D.R., Collapsibility of the upper airway during anesthesia with isoflurane, Anesthesiology 97, 786–793, 2002. 117. D’Honneur, G., Lofaso, F., Drummond, G.B., Rimaniol, J.-M., Aubineau, J.V., Harf, A. and Duvaldestin, P., Susceptibility to upper airway obstruction during partial neuromuscular block, Anesthesiology 88, 371–378, 1998. 118. Eastwood, P.R., Szollosi, I., Platt, P.R. and Hillman, D.R., Comparison of upper airway collapse during general anaesthesia and sleep, Lancet 359, 1207–1209, 2002. 119. Ward, D.S. and Nitti, G.J., The effects of sufentanil on the hemodynamic and respiratory response to exercise, Med. Sci. Sports Exerc. 20, 579–586, 1988. 120. Santiago, T.V., Johnson, J., Riley, D.J. and Edelman, N.H., Effects of morphine on ventilatory response to exercise, J. Appl. Physiol. 47, 112–118, 1979. 121. Eger, E.I., Dolan, W.M., Stevens, W.C., Miller, R.D. and Way, W.L., Surgical stimulation antagonizes the respiratory depression produced by Forane, Anesthesiology 36, 544–549, 1972. 122. Dockery, M.P. and Drummond, G.B., Respiratory response to skin incision during anaesthesia with infusions of propofol and alfentanil, Br. J. Anaesth. 88, 649–652, 2002. 123. Sarton, E., Dahan, A., Teppema, L., Berkenbosch, A., van den Elsen, M. and van Kleef, J., Influence of acute pain induced by activation of cutaneous nociceptors on ventilatory control, Anesthesiology 87, 289–296, 1997. 124. Sarton, E., Dahan, A., Teppema, L., van den Elsen, M.J.L.J., Olofsen, E., Berkenbosch, A. and van Kleef, J., Acute pain and central nervous system arousal do not restore impaired hypoxic ventilatory response during sevoflurane sedation, Anesthesiology 85, 295–303, 1996. 125. Bailey, P.L., Rhondeau, S., Schafer, P.G., Lu, J.K., Timmins, B.S., Foster, W., Pace, N.L. and Stanley, T.H., Dose-response pharmacology of intrathecal morphine in human volunteers, Anesthesiology 79, 49–59, 1993. 126. Petersen, K.L., Jones, B., Segredo, V., Dahl, J.B. and Rowbotham, M.C., Effect of remifentanil on pain and secondary hyperalgesia associated with the heat–capsaicin sensitization model in healthy volunteers, Anesthesiology 94, 15–20, 2001.
5 Response Surface Modeling of Drug Interactions: Model Selection and Multimodel Inference Using the Bootstrap
ERIK OLOFSEN and ALBERT DAHAN Leiden University Medical Center The Netherlands
I.
Introduction
In clinical practice, drugs are often administered in combination to optimize the balance between their therapeutic and adverse effects. Ideally their interaction is synergistic (supra-additive) with respect to the desired effect and only additive or even antagonistic (infra-additive) with respect to their side effects. We studied depression of ventilation and the ventilatory response to hypoxia during combined sevoflurane and alfentanil administration [1] and depression of ventilation and the ventilatory response to hypercapnia during combined propofol and alfentanil administration [2]. To that end, we developed response surface models that capture the relationships between drug concentrations and resulting effects. In the following, we will focus on possible formulations of response surface models and on how to choose between competing models.
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Pharmacodynamic Interaction Models
Pharmacodynamic interaction models attempt to describe the relationship between the concentrations Ci of n simultaneously administered drugs and a resulting measurable effect parameter E under study, e.g., ventilation. If the drugs have the effect of depression from a baseline E0, one can write E ¼ E0 f ðC1 , C2 , . . . , Cn Þ þ 2
ð5:1Þ
where the function f represents the knowledge postulated about the relationship and 2 denotes unexplained variability in measurements of the effect. The function f, visualized in n þ 1-dimensional space, will be referred to as a response surface. A. The Richards Model
The function discussed by Richards [3] attempts to model reaction rates rather than the equilibrium situation, but it may be of value for empirical models. It can be written as
C EðCÞ ¼ E0 1 C50
!!1= 1 1 2
ð5:2Þ
where C50 is the concentration and E ¼ ð1=2ÞE0, and and are shape parameters. When ¼ 1, the equation reduces to the familiar inhibitory sigmoid Emax model EðCÞ ¼ E0
1 1 þ ðC=C50 Þ
ð5:3Þ
while when ¼ 1, it reduces to
C EðCÞ ¼ E0 1 C50
1 2
ð5:4Þ
In the following, the latter model will be referred to as the power model. Parameter C50 may be poorly estimable if it lies outside the concentration range used in the study. It may therefore be better to write 1= C EðCÞ ¼ E0 1 1 l Ch
ð5:5Þ
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where is the fraction of E0 attained when the concentration equals Ch, the midpoint of the concentration range in the study design. An extension for two drugs that respects Loewe additivity (i.e., additivity should hold, irrespective of the values of and , when C1 and C2 both denote concentrations of the same drug [4]) reads as 1= 1= 1= C1 C2 EðC1 , C2 Þ ¼ E0 1 þ 1 1 1 2 Ch,1 Ch,2 ð5:6Þ Its limit when ! 0 equals 1= 1= ! ! C1 1 C2 1 þ log log : 1 2 Ch,1 Ch,2
EðC1 ,C2 Þ ¼ E0 exp
ð5:7Þ B. A Mechanism-based Approach
Suppose that for simultaneous binding of drug molecules to a receptor d½RD ¼ kon ½D ½R koff ½RD dt
ð5:8Þ
where [D], [R] and [RD] are the drug, receptor, and bound receptor concentrations, respectively. For an inhibitory effect model, where effect measure E ranges from E0 to zero, one usually writes ½RT ½RD ½RD ¼ E0 1 E ¼ E0 ½RT ½RT
ð5:9Þ
where [RT ] is the total receptor concentration. Let us instead assume ½RD E ¼ E0 1 ½RX
ð5:10Þ
where [RX ] may possibly be much smaller than [RT ] so that E may be reduced to zero when only a fraction of the receptors is occupied, similar to the concept of receptor reserve. So dE E0 d½RD E0 ¼ ¼ ðkon ½D ½R koff ½RDÞ dt dt ½RX ½RX
ð5:11Þ
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with ½R ¼ ½RT ½RD
and
E ½RD ¼ ½RX 1 E0
ð5:12Þ
which gives dE ½RT ¼ kon ½D E0 ðE0 E Þ þ koff ðE0 E Þ dt ½RX
ð5:13Þ
When [RX ] ¼ [RT ], this reduces to dE ¼ kon ½D E þ koff ðE0 E Þ: dt
ð5:14Þ
In steady-state this yields E ¼ E0
1 , 1 þ ðkon =koff Þ ½D
ð5:15Þ
which is equivalent to Eq. (5.3). However, when ½RX ½RT , ½RT = ½RX E0 ðE0 E Þ, and, by discarding the latter term and incorporation [RT ]/[RX ] in kon, Eq. (5.13) reduces to dE ¼ kon ½D E0 þ koff ðE0 E Þ dt
ð5:16Þ
Note that the interaction between E and D in Eq. (5.13) has disappeared so that the dynamics are closer to those of a classical effect-site model [5], although the presence of in the differential equation still makes them distinct. Furthermore, kon will be dominated by wash-in effects rather than receptor kinetics. In the steady-state we have kon E ¼ E0 1 ½D ð5:17Þ koff which is equivalent to Eq. (5.4). Steady-state relationships between those extremes can be found by introducing a parameter ¼ ½RX =½RT , so that EðCÞ ¼ E0
1 ð1 Þ K ðC=Ch Þ 1 þ K ðC=Ch Þ
with
K¼
1 1 ð1 Þ
ð5:18Þ
where C and denote, like in the previous section, concentration and the fraction of E0 attained when the concentration equals Ch, respectively. C. Modeling Interaction
The current state-of-the-art modeling of interaction is based on two fundamental ideas [6]. First, the combination of two drugs is considered
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to act like a single drug with a certain concentration–effect relationship. Second, the properties of this virtual drug, and therefore the parameters of its concentration–effect relationship, are only dependent on the ratio of the concentrations of the two drugs, given by Q ¼ U1/(U1 þ U2), where U1 ¼ C1/Ch,1 and U2 ¼ C2/Ch,2. In the studies described below, interaction was incorporated by writing 1= 1= 1= C1 C2 EðC1 ,C2 Þ ¼ E0 1 þ IðQÞ 1 1 1 2 Ch,1 Ch,2 ð5:19Þ where I(Q) is a spline function with parameters Imax and Qmax, which denote the maximum value of the interaction and the value of Q at which I(Qmax) ¼ Imax, respectively. For further details, see Ref. [1]. III.
Model Selection and Multimodel Inference
In the previous section, we discussed two competing model formulations that differently link the same two extremes: the power model (Eq. (5.19) with ¼ 1) and the inhibitory sigmoid Emax model (Eq. (5.19) with ¼ 1). In order to determine which of those two models is best suited for our experimental data, we need an appropriate criterion of what is best. We will now briefly introduce the likelihood ratio test and Akaike’s information-theoretic criterion (AIC); for a complete discussion, see Ref. [7]. The likelihood ratio test is based on the statistic C2 ¼ 2 ðLLr LLf Þ
ð5:20Þ
where LLr and LLf are the maximized log-likelihoods of the reduced and full models, respectively, C2 is approximately 2 distributed with q degrees of freedom and q is the number of fixed parameters in the reduced model. It can be used to test the hypothesis that a parameter has a certain fixed value. However, the -level of the test is rather arbitrary, especially for a chain of nested models, and this criterion cannot be used for nonnested models. Alternatives are Akaike’s information-theoretic criterion (AIC) and its variant (AICc) with a correction for small sample sizes, which are given by AIC ¼ 2LL þ 2k AICc ¼ 2LL þ 2k
N Nk1
ð5:21Þ ð5:22Þ
where LL is the maximized log-likelihood of the model under consideration, k is the number of adjustable parameters and N is the number of samples.
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AIC is an estimate of (2 times) the relative information loss when truth is approximated by a model; therefore the model with the lowest AIC can be considered to be best. As truth is unknown, only relative values can be evaluated. The bias (equal to k) originates from the fact that the parameters of the model have to be estimated. It may well be that alternative models are almost equally suited to capture the information in the data and discarding them may result in biased and incorrectly precise parameter estimates and inferences. Models can be assigned [7] so-called Akaike weights which are given by expðð1=2Þi Þ w i ¼ PR r¼1 expðð1=2Þr Þ
ð5:23Þ
where R is the number of models, and i the Akaike differences i ¼ AICi AICmin
ð5:24Þ
and AICmin is the value of AIC for the best model (note that the wi do not depend on it). When parameters have equivalent interpretations across models, the weights can be used to obtain model-averaged estimates of parameters and their variances; otherwise the predicted expected response variables from each model may be averaged. When more than one data set is available, model weights can also be obtained by model selection frequencies which are given by the number of times a particular model is chosen for each data set. In the following example, multiple data sets are generated by means of Monte Carlo simulation. A. An Example
To illustrate the use of AIC, we now focus on the following simple model: yi ¼ a þ 2 i
ð5:25Þ
where yi ði ¼ 1, . . . , NÞ are samples, a is a constant and 2i are independent Gaussian variates with variance 2 : The maximum likelihood estimators of the parameters are well known, and given by a^ ¼
N 1X yi N i¼1
and
^ 2 ¼
N 1X ðyi a^ Þ2 N i¼1
ð5:26Þ
The question now arises whether the data support estimation of parameter a; it could be better to fix it at some value,pfor ffiffiffiffi example zero, which simplifies the model. From a^ and VARða^ Þ ¼ ^ 2 = N, a confidence interval can be constructed to test the hypothesis H0: a ¼ 0. However, we will proceed in
Response Surface Modeling of Drug Interactions
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A
B 0.6 Selection frequency
Selection frequency
0.3
0.2
0.1
0
0.5 0.4 0.3 0.2 0.1 0
-2
-1
0
1 a
2
3
4
-2
0
1 a
2
3
4
C
0.8 Selection frequency
-1
0.6 0.4 0.2 0 -2
-1
0
1 a
2
3
4
Figure 5.1 Normalized selection frequencies for the illustrative model given by Eq. (5.25) using data from Monte Carlo simulations. A. Model set consisted of seven alternatives with fixed values of a. B. Model set consisted of M0 with fixed value of a ¼ 0 and M1 where a was adjustable. C. Model set consisted M0 with fixed value of a ¼ 1 and M1 where a was adjustable.
a different way, since in practice we can be sure that H0 is false, and our aim is to minimize the loss of information that occurs when the data are approximated by a model. Monte Carlo simulation is a general method for assessing the properties of estimators [8]. From the model, B ¼ 1,000 realizations were generated, with a ¼ 1, 2 ¼ 25, and N ¼ 25. Seven models were fitted to the data using NONMEM [9]. Parameter a was fixed to each of the values 2, 1, . . . , 4; 2 remained adjustable. The Akaike criterion was used to select the best model for each of the B realizations. Figure 5.1A shows the fraction of times each model was selected. The probability of selecting the model with a ¼ 0 was 0.241. This value corresponds nicely to the probability of 0.242 that a^ is between (0.5, 0.5) obtained from its theoretical distribution function (although this relationship is not necessarily exact).
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The model with a fixed value of a ¼ 1 is best. However, in practice, the value of a is unknown and the models with alternative values of a also have considerable weight. A more often used model set would consist of M0 where a is fixed to zero, together with M1 where a is adjustable. Figure 5.1B shows the probability density function of model selection and the corresponding value of a, estimated from 10,000 realizations. The best model was M0 with selection probability 0.6955. Again, alternative values of a have considerable weight. A possibility [7] here is to calculate a weighted value, namely a^ ¼ 0:6955 0 þ 0:3045 mean of alternative estimates of að2:044Þ 0:6225: So this procedure removes part of the bias caused by considering only the best model (a ¼ 0, while true a ¼ 1). It is also possible [7] to calculate a model-averaged value of the variance of a^ , since the variance of the fixed a ¼ 0, being zero, is surely underestimated. In this example, VARfa^ g 0:9783: Notice that the hypothesis H0: a ¼ 0 would not have been rejected, while the single model with fixed a ¼ 0 is not optimal. We could, by accident, have chosen M0 with fixed a ¼ 1. By using the procedure produced above, we find that the probability that AIC selects this model is approximately 0.8780, a^ 1:008 and VARfa^ g 0:070 (Figure 5.1C). Of course, the model set must be carefully defined a priori; it is statistically incorrect to locate a fixed value of a of M0 that minimizes its variance after the data have been collected. When it has to be located, it should be a free parameter or else many fixed values could be tried as the distribution of the model weights is approximately equivalent to the one of a^ if a were free (Figure 5.1A). There should be no variability in the way a fixed value of a is chosen; eliminating a parameter by fixing it to zero is a general way to simplify a model. It is clear that it is not appropriate to consider only the best model (with a ¼ 1) as this gives a false sense of precision. The larger the standard error of a when it is free, the more likely it can be fixed, as the data do not provide information about its value. When there is an estimate available from another study, this estimate could be incorporated using the Bayesian approach or again fixed if the uncertainty is negligible, such as for fundamental constants from physics. In practice, there is usually only one data set and model selection frequencies are obviously not available. Model selection or averaging could be done using the Akaike weights as defined above, but we will now consider estimating model selection frequencies by a method called the bootstrap. IV.
The Bootstrap
A. Introduction
The bootstrap is an alternative method for assessing the properties of estimators [10]. To explain the principle underlying the bootstrap, consider
Response Surface Modeling of Drug Interactions
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the definition of the mean of a discrete probability distribution pk with K possible events: ¼
K X
k pk
ð5:27Þ
k¼1
In practice, the pk are unknown, but they can be estimated by the number of times k is observed divided by the total number of observations N, by assigning pi ¼ 1/N to each observation xi. Therefore can be estimated by the well-known formula ¼
N 1X xi N i¼1
ð5:28Þ
So the true distribution F (discrete or continuous) is approximated by an empirical distribution F^ consisting of weights 1/N at the observations, which can be plugged-in to obtain estimates of its parameters. As the observations xi are samples from F^, one realizes that repeating an experiment (for example to construct confidence intervals) can be simulated by resampling, with replacement, from F^: The estimator under study is applied to each replicate data set and hence its bias and variance can be assessed. This is important in particular when other means of estimating them are lacking, such as the properties of the median. The fact that this is possible may be surprising, but actually it is equally surprising that standard errors can be obtained from a maximum likelihood estimator which is also based on a single data set. Maximum likelihood estimators do have welldefined properties, but in general only asymptotically (so when the number of data points is infinite); the bootstrap can then be used to study them when the amount of data is small. But in the following, we will study bootstrapaided model selection in particular. B. An Example
We go back to the simple example of the previous section and study how the bootstrap can be employed to guide model selection. One data set was generated from the model with parameters given earlier. From this one data set, B ¼ 1,000 replicates were generated by sampling with replacement such that the number of samples remained N ¼ 25. To each replicate, the seven models were fitted and the fraction of times each model was selected according to the Akaike criterion is shown in Figure 5.2A. This distribution resembles the Normal distribution of a^ with a^ 0:468 and ^ 2 1:16 (these estimates are obtained from the single data set used for the bootstrap). Figure 5.2B and Figure 5.2C show the probability density functions of model selection and the corresponding value of a, estimated
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Selection frequency
Selection frequency
0.3
A
0.3
0.2
0.1
B
0.6 0.4 0.2 0
0 -2
-1
0
1 a
2
3
4
-2
0
1 a
2
3
4
C
0.5 Selection frequency
-1
0.4 0.3 0.2 0.1 0 -2
-1
0
1 a
2
3
4
Figure 5.2 Normalized selection frequencies for the illustrative model given by Eq. (5.25) using bootstrap data sets. A. Model set consisted of seven alternatives with fixed values of a. B. Model set consisted of M0 with fixed value of a ¼ 0 and M1 where a was adjustable. C. Model set consisted M0 with fixed value of a ¼ 1 and M1 where a was adjustable.
from 10,000 realizations with the previously defined model sets. It is important here to forget that the true value of a ¼ 1 because in most life science experiments, the true values are unknown, and reality is always more complicated than the models under consideration. The best we can do is find a good approximation of the information contained in the data, and ensure that parameter estimates are associated with a proper measure of their uncertainties. In this example, we could select model M0 with confidence factor 0.8371, which differs from 0.6955 as obtained from the Monte Carlo simulations. However, this factor itself is a random variable and varies between experiments (just as a confidence interval does). The point is that the bootstrap enables us to obtain a measure of model selection uncertainty. Model selection frequencies are similar, but in general not equal to, the Akaike weights. In this example, the Akaike weight of model M0 ¼ 0.7480
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and model M1 has associated a^ ¼ 0:468: Further research is needed to assess their respective properties. However, a practical advantage of using the Akaike weights is that no computer-intensive bootstrap runs are needed. V.
Applications
A. Interaction between Sevoflurane and Alfentanil
Nine healthy male volunteers participated, after approval of the protocol by the local Human Ethics Committee, in a study designed to assess the effects of sevoflurane and alfentanil, given separately and in combination, on resting normoxic ventilation and on ventilation due to acute isocapnic hypoxia (duration of hypoxia ¼ 3 min, PETO2 ¼ 48 mmHg). The end-tidal concentrations of sevoflurane (Csev) used were 0, 0.1, 0.2 and 0.3%; the target alfentanil plasma concentrations (Calf) used were 0, 10, 20, 30, 40, and 50 ng/ml. The nature of the interaction was assessed for, among other things, the steady-state ventilation ðV_ i Þ and its dependence of the arterial blood saturation ðV_ i =Sp O2 Þ: For further details, see Ref. [1]. A population analysis using NONMEM [9] yielded a linear relationship between both sevoflurance and alfentanil concentration and (hyperisocapnic, normoxic) ventilation with synergistic interaction between sevoflurane and alfentanil. Parameter estimates were associated with a relatively large standard error and since C50 cannot be negative, the distributions were most probably skewed. We employed the bootstrap to assess these parameter distributions and confidence intervals (using the bias-corrected and accelerated (BCa) method [10]) from B ¼ 1,000 replicate data sets. Histograms of the parameter estimates are shown in Figure 5.3. The peaks in the bins at the maximum x-value correspond to quite a number of large parameter estimates and are another sign of their skewed distributions. The 50% and 95% confidence intervals were: C50, alf 62.8–92.2 and 45.9–360 ng/ml; C50, sev 1.14–2.81 and 0.567–308%; Imax 1.71–2.24 and 1.38–6.66. In particular, the C50 of sevoflurane was poorly determined form the data as the concentration range used in the study (0–0.4%) was well below the estimated C50. In Ref. [2], we found that transforming the model parameters from C50 to as described in Section II considerably improved parameter estimation. With this model, parameter Qmax could be fixed to 0.5, which means that interaction is maximal at the diagonal that crosses the concentration plane designed for the experiment, and is therefore not a coincidental value. Histograms of parameter estimates using the transformed model with fixed ¼ 1, fixed Q max ¼ 0.5 and free Imax are shown in Figure 5.4. The 50% and 95% confidence intervals
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Olofsen and Dahan A
0.2
0.1 P (C50, sev)
0.15
P (C50, alf)
B
0.125
0.1
0.05
0.075 0.05 0.025
0
0 0
50
100 150 C50, alf
0
200
1
2 3 C50, sev
4
C
0.25 P (Imax)
0.2 0.15 0.1 0.05 0 0
1
2
3 Imax
4
5
Figure 5.3 Modeling depression of ventilation by sevoflurane and alfentanil: histograms for the estimates of parameters C50,alf (A), C50,sev (B), and Imax (C).
were: alf 0.738–0.812 and 0.657–0.890; sev 0.849–0.922 and 0.757–0.971; Imax 1.27–1.52 and 0.969–1.97. The value of maximum interaction Imax was determined more precisely, but was also closer to 1. Furthermore, we wondered which model would best fit the data: the inhibitory sigmoid Emax model or the power model described in Section II. The bootstrap model selection frequencies in Table 5.1 provide answers to both questions: the information in the data was best captured by the sigmoid Emax model with interaction. Rather than discarding the alternative models, we keep all eight of them, but with weight factors determined by the model selection frequencies. Averaging model parameters across the different models is not meaningful here as some have different typical values. However, it is possible to obtain mode-averaged predictions, using the selected models and their parameters from each replicate data set, and to determine their median prediction, and 50%, 80%, and 90% prediction intervals as shown in Figure 5.5. It shows prediction intervals for the single drugs and for their combination when interaction is maximal (at Q ¼ Qmax). Note that the concentrations extend those used in the experiments by 50% so the intervals
Response Surface Modeling of Drug Interactions
145 B
0.15
0.15
0.1
0.1
P (λsev)
P (λalf)
A
0.05
0.05
0
0 0.5
0.625
0.75 λalf
0.875
0.5
1
0.625
0.75 λsev
0.875
1
C 0.25
P (Imax)
0.2 0.15 0.1 0.05 0 0
1
2
3
Imax
Figure 5.4 Modeling depression of ventilation by sevoflurane and alfentanil: histograms for the estimators of parameters alf ðAÞ, sev ðBÞ and Imax (C). Table 5.1 Depression of ventilation by sevoflurane and alfentanil: power and sigmoid Emax model selection frequencies obtained from 1,000 bootstrap data sets. The linear model is the power model with ¼ 1; additive interaction is present when Imax ¼ 1
¼ 1; 6¼ 1; ¼ 1; 6¼ 1;
Imax Imax Imax Imax
¼1 ¼1 6¼ 1 6¼ 1
Power Model
Sigmoid Model
0.113 0.012 0.193 0.003
0.029 0.025 0.349 0.275
plotted in that range are extrapolations. Furthermore, the drug effects were normalized to baseline. Sevoflurane was less potent than alfentanil in depressing ventilation, also with respect to their interaction, since adding sevoflurane (going from the top left to the top right panel) has almost no
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effect, while adding alfentanil (going from the bottom to the top right panel) has a large effect. Parameter V_ i =Sp O2 , the hypoxic ventilatory response, was found to be linearly dependent on alfentanil and sevoflurane concentration, without interaction. In Figure 5.6 histograms of parameter estimates are shown from a bootstrap analysis with fixed ¼ 1, fixed Qmax ¼ 0.5 and free Imax. Parameter Imax was estimated to be very close to 1. Model selection frequencies are given in Table 5.2. In this case the linear, additive interaction model was best. The multimodel median response surface is shown in Figure 5.7; the prediction intervals are shown in Figure 5.8. The median curves show remarkable linearity, but the top curves (with smallest depression) show some sigmoidicity. The curvature at low concentrations
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Table 5.2 Depression of the hypoxic ventilatory response (Vi/SpO2) by sevoflurane and alfentanil: power and sigmoid Emax model selection frequencies obtained from 1,000 bootstrap data sets. The linear model is the power model with ¼ 1; additive interaction is present when Imax ¼ 1
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cannot be captured by the linear model. However, the sigmoid Emax model is a symmetric curve so that a curvature at low concentrations is always mirrored at high concentrations. The question now is what would happen at concentrations higher than those used in the experiments. It is most probably not true physiologically that parameter V_ i =Sp O2 will reach zero only at infinite drug concentrations. Furthermore, predictions with negative values should be admissible since the depressant effects of central hypoxia may more than counteract the effects of hypoxia at the peripheral chemoreceptors [1,11]. Multimodel inference allows us to deal with these possibilities and avoids falsely precise, and therefore less accurate, predictions.
B. Interaction between Propofol and Remifentanil
In another study, 22 healthy male volunteers participated after approval of the protocol by the local Human Ethics Committee. They received targetcontrolled infusions of propofol and remifentanil such that at least four concentration combinations were achieved. The target concentrations were chosen at evenly spaced design points between 0 and 2 ng/ml remifentanil
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and 0 and 2 mg=ml propofol. At these drug levels, ventilation and the ventilatory response to hypercapnia were measured. Response surfaces _ i and ventilation at a fixed end-tidal PCO of for resting ventilation V 2 55 mmHg V_ i,55 were constructed using a population analysis with NONMEM [9]. For further details, see Ref. [2]. In that study, we found that both V_ i and V_ i,55 were best described by the power model with free , fixed Qmax ¼ 0.5 and free Imax. Model selection frequencies (with free and fixed Qmax ¼ 0.5; number of replicate data sets B ¼ 1,000) are given in Table 5.3 for both V_ i and V_ i, 55 : Clearly, the power model with interaction is best, especially for V_ i,55 : As parameter distributions were approximately normal, showing their histrograms would not provide much information
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Table 5.3 Depression of ventilation (Vi) and ventilation at 55 mmHg (Vi,55) by propofol and remifentanil: power and sigmoid Emax model selection frequencies obtained from 1,000 bootstrap data sets. Additive interaction is present at Imax ¼ 1 Power Model
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beyond the standard errors. Baseline V_ i and V_ i,55 of the best models were 9.4 0.3 (SE) and 31.4 1.5 (SE), respectively. The multimodel median response surface for V_ i,55 obtained by the bootstrap model selections is shown in Figure 5.9; prediction intervals are shown in Figure 5.10 for V_ i and Figure 5.11 for V_ i,55 . The curves for V_ i,55 show a rapid decrease of the respiratory controller’s sensitivity to CO2 and subsequently follow the depression of baseline ventilation. Parameter Imax was 1.9 0.2 (SE) for V_ i and 1.2 0.1 (SE) for V_ i,55 of the best models. Propofol was less potent in depressing V_ i than V_ i,55, but the larger value of Imax partly compensates for this. Their interaction is such that adding propofol (going from the top left
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to the top right panels) has a small effect, while adding remifentanil (going from the bottom to the top right panels) has a large effect. VI.
Conclusions
Respiratory depression by combined administration of sevoflurane and alfentanil, or propofol and remifentanil, was described by response surface modeling. The adequacy of competing model formulations was assessed by a bootstrap-aided model selection strategy based on AIC. The bootstrap allowed us both to assess model selection stability and to obtain multimodel inferences such as predicting the balance between therapeutic and adverse effects. Multimodel inference
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avoids biased and incorrectly precise predictions based on a single, possibly marginally better, model. However, from our analyses we do conclude that the power model is more suitable to describe respiratory depression that the classical sigmoid Emax model, especially if we want to take loss of ventilatory stability and apnea into account. References 1.
Dahan, A., Nieuwenhuijs, D.J.F., Olofsen, E., Sarton, E.Y., Romberg, R.R. and Teppema, L.J., Response surface modeling of alfentanil–sevoflurane interaction on cardiorespiratory control and bispectral index, Anesthesiology 94, 982–991, 2001.
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6.
7. 8. 9. 10. 11.
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Nieuwenhuijs, D.J.F., Olofsen, E., Romberg, R.R., Sarton, E.Y., Ward, D.S., Engbers, F.H.M., Vuyk, J., Mooren, R.A.G., Teppema, L.J. and Dahan, A., Response surface modeling of remifentanil–propofol interaction on cardiorespiratory control and bispectral index, Anesthesiology, 98, 312–322, 2003. Richards, F.J., A flexible growth function for empirical use, J. Exp. Botany 10, 290–300, 1959. Berenbaum, M.C., What is synergy? Pharm. Rev. 41, 93–141, 1989. Sheiner, L.B., Stanski, D.R., Vozeh, S., Miller, R.D. and Ham, J., Simultaneous modeling of pharmacokinetics and pharmacodynamics: application to d-tubocurarine, Clin. Pharm. Ther. 24, 358–371, 1979. Minto, C.F., Schnider, T.W., Short, T.G., Gregg, K.M., Gentilini, A. and Shafer, S.L., Response surface model for anesthetic drug interactions, Anesthesiology 92, 1603–1616, 2000. Burnham, K.P. and Anderson, D.R., Model Selection and Multimodel Inference, 2nd edn., New York, Springer, 2002. Kotz, S. and Johnson, N.L., Encyclopedia of Statistical Science, Vol. 5., New York, Wiley, 1985. Beal, S.L. and Sheiner, L.B., NONMEM User’s Guides, Nonmen Project Group, University of California at San Francisco, San Francisco, 1999. Efron, B. and Tibshirani, R.J., An Introduction to the Bootstrap, New York, Chapman and Hall, 1993. Sarton, E.Y. and Dahan, A., Sites of respiratory action of opioids, in On the Study and Practice of Intravenous Anaesthetics, Vuyk, J., Engbers, F.H.M. and Groen-Mulder, S.M., eds., Dordrecht, The Netherlands, Kluwer Academic Publishers, 2000.
6 Respiratory Neuroplasticity: Respiratory Gases, Development, and Spinal Injury
DAVID D. FULLER and GORDON S. MITCHELL University of Wisconsin Madison, Wisconsin
RYAN W. BAVIS Assistant Professor of Biology Bates College Lewiston, Maine
I.
Introduction
Since prolonged failure of the respiratory control sysem is not compatible with life, the mechanisms underlying respiratory control must be robust under a wide range of conditions. However, a robust neural system need not be a rigid circuit. Neuroplasticity enables appropriate adaptations to frequent or chronic disturbances that initiate active modifications in respiratory control such as weight gain or loss, altitude exposure or injury [1]. Studies of respiratory plasticity are intended to reveal how and why experience or changing conditions influence the control of breathing. Further, the respiratory system provides an ideal model to explore fundamental mechanisms of neuroplasticity for at least two reasons. First, the respiratory neural control networks produce a spontaneous rhythmic and quantifiable motor output under a wide range of in vivo and in vitro conditions. Second, clear functional significance can be ascribed to respiratory motor output: it represents breathing. Many influential models of neuroplasticity (e.g., hippocampal long-term potentiation; [2,3]) lack these advantages. 155
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We have begun to make progress toward understanding the mechanisms and manifestations of respiratory neuroplasticity [1,4,5]. In this chapter, we will review the literature concerning three forms of respiratory plasticity including plasticity induced by altered respiratory gases in adults, plasticity specific to development, and plasticity following injuries to the spinal cord. These topics illustrate diverse mechanisms and functional significance of plasticity in respiratory control. Other interesting and important examples of respiratory plasticity are omitted to allow a more comprehensive review of these topics, which represent the most thoroughly studied aspects of respiratory neuroplasticity to date. The reader is also referred to a recent series of review articles on respiratory neuroplasticity [1,6–10] . These articles provide an in-depth overview, and illustrate the emerging interest and excitement regarding this topic. A. Plasticity vs. Modulation
Plasticity and modulation are related but distinct properties of neural systems [1]. These properties are not mutually exclusive, which may lead to confusion. For example, modulation of a neural process may induce or maintain altered system characteristics, thereby eliciting plasticity. To promote effective communication among respiratory physiologists, Mitchell and Johnson [1] proposed the following working definitions of plasticity and modulation pertaining to respiratory motor control: Plasticity: ‘‘a persistent change in the neural control system (morphology and/or function) based on prior experience.’’ Modulation: ‘a neurochemically induced alteration in synaptic strength or cellular properties, adjusting or even transforming neural network function.’ In other words, respiratory plasticity describes an effect that outlasts the stimulus for a period of seconds to years, whereas neuromodulation generally occurs on a relatively short time scale (e.g., seconds to minutes) and is rapidly reversed when the neuromodulator is no longer present and functionally active. Plasticity and modulation are not static processes; they are susceptible to modification by prevailing conditions or experience [11–13]. Metaplasticity (i.e., plastic plasticity) describes a change in the expression of plasticity due to experience [11,12,14]. Similarly, metamodulation refers to modulation of a modulatory process [1,13]. B. Where does Respiratory Neuroplasticity Occur?
The neurons that control breathing represent a complex, integrative network (Figure 6.1). Redundancy in respiratory control mechanisms is evident in the mammalian central nervous system (CNS) [7,15,16], and
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Figure 6.1 A. Potential sites of plasticity in respiratory motor control. Alterations in respiratory motor output could reflect neuroplasticity at multiple sites including: (1) peripheral chemoreceptors; (2) the nucleus of the solitary tract; (3) the ventral respiratory group, including the pre-Bo¨tzinger complex; (4) supramedullary structures including the cerebral cortex, thalamus and cerebellum; (5) afferent inputs to the spinal cord; (6) respiratory motoneurons or local interneurons (see Figure 6.1B); or (7) the neuromuscular junction. Each of these possibilities is discussed in the text. B. Potential cellular and synaptic mechanisms underlying respiratory motor plasticity. Respiratory neuroplasticity at any site may result from the cellular/synaptic mechanisms exemplified here for a respiratory neuron located in the spinal cord. Plasticity may involve multiple mechanisms including: (1) changes in neuron properties such as somal size and membrane properties (e.g., resistance, rheobase current, etc.); (2) changes in synaptic efficacy induced by neuromodulators such as serotonin (see Figure 6.2); (3) alterations in synaptic efficacy due to prior activity within that synapse (i.e., activity-dependent synaptic plasticity); (4) revelation of pre-existing synaptic connections that were previously ineffective (i.e., silent synapses); (5) alterations in neuromodulator function due to changes in the neuromodulator (i.e., metaplasticity) or in release, reuptake and/or receptor activation; (6) growth of new synapses (excitatory or inhibitory) or changes in arborization of existing connections (i.e., sprouting).
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this redundancy has a physiological advantage: failure in a single component will not necessarily cripple the system. Similarly, plasticity in respiratory control need not be restricted to a discrete location. Sites suspected to contribute to different models of respiratory plasticity include peripheral chemoreceptors or chemosensory neurons [6,17–19], the nucleus of the solitary tract [20–22], pre-motoneurons, modulatory neurons (e.g., serotonergic or noradrenergic; [1,10,23]), pontine respiratory neurons [24,25], brainstem rhythm-generating neurons [26,27], spinal neurons ([28–30]; Figure 6.1), and the neuromuscular junction [9,31]. Plasticity rostral to the brainstem may also impact respiratory control as shown by classical and operant conditioning studies [32–36]. Figure 6.1 presents a simplified scheme for respiratory control and identifies potential sites of respiratory neuroplasticity; presumably plasticity can occur at any level in the network, and possibly many levels at once. II.
Plasticity Induced by Respiratory Gases in Adult Mammals
Changes in respiratory gases and respiratory plasticity are of particular interest for several reasons. First, it is common for humans to experience alterations in blood gases, particularly during pulmonary disease, sleepdisordered breathing or ascent to altitude. In healthy individuals, plasticity may optimize or smooth respiratory motor output during or following alterations in respiratory gases. A failure of plasticity may contribute to disorders such as obstructive sleep apnea (OSA) or sudden infant death syndrome (SIDS). Here we review the ventilatory response during and following acute and chronic exposure to both sustained and intermittent alterations in inspired oxygen and carbon dioxide. Acute has been defined as one hour or less [37]. The distinction between sustained and intermittent exposures is important since it has become clear that the specific pattern of respiratory stimulation can be an important determinant of the forms of plasticity that will be induced [4,38]. A. Hypoxia
Reductions in arterial oxygen tension activate a complex feedback system that typically increases alveolar ventilation in adults. The primary source for reflex ventilatory stimulation during hypoxia is the carotid body [37]. Hypoxia also stimulates chemosensitive cells in the aortic bodies [39,40] and some neurons in the brainstem [41–43], although their contributions to breathing under normal circumstances are either minimal or not understood. However, their minimal role in normal conditions does not rule out increased contributions to the control of breathing due to
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plasticity. For example, the aortic chemoreceptors become increasingly important following carotid denervation, particularly in young animals [7,44,45]. The nucleus of the solitary tract in the medulla provides the initial central synaptic relay for peripheral chemoafferent neurons. Subsequent projections include the ventrolateral medulla and midline raphe neurons [46], ultimately reaching respiratory motoneurons or other CNS regions including the cerebellum [47,48]. Acute Hypoxia (51 h)
The acute hypoxic ventilatory response is characterized by a hyperbolic increase in minute ventilation as the partial pressure of arterial oxygen (PaO2) declines [37,49]. However, even during isocapnic hypoxia, the temporal pattern of breathing (i.e., changes in inspiratory tidal volume vs. frequency) is variable within and among species [37,49]. Certain neurochemicals that contribute to the acute hypoxic response have been reviewed in detail [21,50]. Following the acute increase in ventilation, hypoxia evokes several time-dependent changes in respiratory motor output including short-term potentiation (STP), short-term depression (STD), and hypoxic ventilatory depression (HVD). These responses represent discrete, time-dependent mechanisms that depend on the severity, duration and pattern of hypoxia [37,49,51]. One consideration is whether the underlying mechanisms represent modulation or plasticity. For example, STP (see below) occurs within seconds to minutes of the initial acute response. Increases in ventilation during STP could represent plasticity initiated during the early stages of chemoafferent input. Alternatively, ongoing modulation of chemoafferent stimulation could also produce STP. Resolution will require a more thorough understanding of the mechanisms underlying these respiratory responses. Short-term potentiation (STP)
Slow (second to minute) increases in ventilation following the initial, abrupt increase in ventilation during hypoxia are known as STP [49]. Following hypoxia, the slow decline of respiratory motor output towards pre-hypoxia baseline values may represent decay of the STP mechanism [49,51]. However, the onset of STP is more rapid than the decay in ventilation following hypoxia [52], which may indicate that the decay in ventilation occurs via a distinct mechanism. STP has been demonstrated in awake humans [53,54], goats [55], ducks [56], and mice [57–59], and anesthetized cats [52], dogs [28,60], and rats [61,62]. It is usually expressed as an increase in tidal volume or respiratory neurogram amplitude (vs. frequency; [49]) after the acute hypoxic response. The physiological significance of STP is unknown, but this mechanism may ‘smooth’ ventilation and promote
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stability of respiratory motor output during and following hypoxia [51,63]. Failure of the STP mechanism is associated with pathologies including OSA and congestive heart failure since neither patient population exhibits STP during or following hypoxia [64,65]. STP can be evoked by electrical stimulation of carotid chemoafferent neurons, indicating that it results from a central neural mechanism [20,51,61]. The manifestation of STP is modified by vagal inputs [60], the magnitude and duration of the hypoxic response [66,66a], and prevailing CO2 levels [55,67]. STP may be genetically determined as it is absent in certain mouse strains [57]. Several laboratories have attempted to identify molecules necessary for the expression of STP. Millhorn et al. [68] showed that serotonergic, dopaminergic, and noradrenergic drugs do not affect STP in anesthetized cats. Since nitric oxide (NO) is important for other forms of plasticity, several investigators have hypothesized that it plays a role in STP [21,58,59]. STP is not seen in mice treated with a nitric oxide synthase-1 (NOS-1) inhibitor [69] or in mutant mice deficient in NOS-1 [69]. Kline et al. [69] suggest that NO alters ionic conductances in central respiratory neurons during hypoxia, resulting in STP. Further, NO may be involved in STP through the formation of nitrosothiols [70]. An N-methyl-D-aspartate (NMDA) receptor involvement in STP is suggested by observations that STP is attenuated by NMDA receptor antagonists in anesthetized rats, although the location of the relevant receptors is uncertain [62,63,71]. Other postulated but unproven mechanisms of respiratory STP include calcium accumulation in premotoneurons [52] and substance P release in the nucleus tractus solitarius (NTS) [20]. Phrenic STP may reflect a spinal mechanism, possibly within bulbospinal synaptic connections to spinal phrenic motoneurons [28]. At least three experiments have produced data consistent with this hypothesis [28,29,72]. McCrimmon and colleagues [28] showed that descending synaptic inputs to phrenic motoneurons can be potentiated via NMDA receptor-dependent mechanisms. A short latency (1–2 ms) phrenic compound action potential was uniquely revealed following paired pulse spinal cord stimulation. The new potential was eliminated after systemic administration of the NMDA antagonist MK-801, consistent with the NMDA dependence of STP [62]. A form of STP is induced by high frequency (100 Hz) stimulation of the C1–C2 lateral funiculus in anesthetized rats [29]. Intracellular recordings indicate that short-lasting phrenic motoneuron depolarization and enhanced excitatory post-synaptic potentials (EPSP) occur coincident with STP in this model, thereby providing strong evidence for a spinal site of action [29]. Potentiation of spinal inputs to respiratory motoneurons has also been demonstrated in an in vitro turtle preparation [72]. High-frequency electrical stimulation of spinal synaptic inputs to respiratory motoneurons caused a persistent potentiation (minutes) of
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evoked potential amplitude [72]; the short latency of the evoked potentials (5 1 ms) indicates that spinal synaptic transmission was enhanced. The studies discussed above demonstrate a spinal effect similar to STP, but currently it is unknown if similar mechanisms contribute to STP during and following hypoxia. Nevertheless, these data strongly suggest that plasticity in or around respiratory motoneurons may contribute to STP. For example, potentiation following spinal stimulation [28,29,72,73] could result from plasticity within motoneurons, interneurons, or pre-synaptic inputs (all of which could reflect a spinal mechanism). However, STP of supraspinal motor output (e.g., hypoglossal) could occur by a similar mechanism. Thus, one possibility is that STP arises within discrete motor nuclei (e.g., phrenic, hypoglossal), and does not reflect an overall increase in descending respiratory drive. This hypothesis is supported by studies showing that phrenic and hypoglossal STP can be dissociated in anesthetized cats [74]. Specifically, stimulation of the carotid sinus nerve (CSN) elicits STP in both phrenic and hypoglossal motor output, whereas superior laryngeal nerve stimulation evoked STP of hypoglossal, but not phrenic, motor output. Since the hypoglossal motor output was uniquely affected following superior laryngeal nerve stimulation, STP must be induced independently in different respiratory motor outputs, although the cellular/synaptic mechanisms may be the same. Hypoxic ventilatory depression (HVD) or ‘roll-off’
When hypoxia is sustained for minutes to hours, HVD develops (also known as hypoxic ventilatory roll-off; [37,75]). In adult humans, steadystate isocapnic hypoxia induces an initial acute increase in ventilation that peaks after approximately 3–5 min. This peak is followed by a ‘roll-off’ to a new, lower ventilation that is subsequently maintained [37,75]. As reviewed by Bisgard and Forster [75], potential mechanisms underlying HVD include (1) altered pulmonary mechanics; (2) reduced peripheral (carotid body) chemosensitivity; (3) a direct effect of carotid chemoafferent neurons on medullary excitability; (4) cerebral hypocapnia and/or alkalosis; (5) direct effects of hypoxia on the CNS (i.e., hypoxic brain depression), or (6) reduction in metabolic rate during hypoxia. The mechanism(s) of HVD have not yet been resolved, but there is general agreement that changes in pulmonary mechanics, altered metabolic rate or hypocapnia cannot explain HVD in most circumstances [37,49]. Furthermore, most experimental evidence suggests that a reduction in peripheral chemosensitivity cannot explain HVD, at least in adult animals [37]. The preponderance of evidence indicates that HVD results from a central neural mechanism activated by input from peripheral chemoreceptors. The neurochemicals required for HVD remain under investigation. Serotonin and/or opioids are not necessary for HVD [37]. However adenosine, dopamine, GABA, protein kinase C (PKC), and platelet-derived
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growth factor b (PDGF-b) are strong candidate molecules. An adenosine receptor antagonist (aminophylline) attenuates HVD in humans [76]. However, another adenosine receptor antagonist, theophylline, does not prevent HVD in goats [77]. Dopamine may be involved in HVD because the dopamine antagonist haloperidol blocks HVD in cats [78]. A build-up of the inhibitory neurotransmitter GABA has also been implicated in HVD [37]. The decline in phrenic motor output during 20 min of hypoxia (10% O2) corresponds to a gradual increase in GABA in the ventrolateral medulla [79]. Gozal and Gozal [80] showed that a ventilatory decline occurs in parallel to reductions in NTS PKC activity in hypoxic rats. Because PKC may facilitate respiratory motor output during hypoxia, it was suggested that reductions in PKC may contribute to HVD. Based on known links among PKC, NMDA receptors, and PDGF-b, Gozal and colleagues hypothesized that hypoxia triggers PDGF-b release in the brainstem, thereby contributing to HVD. They showed that pharmacological inhibition of PDGF-b receptors abolishes HVD in mice [80a]. The overall picture that emerges is that HVD may involve a complex interaction between multiple neurochemicals, and may reflect several distinct mechanisms. Short-term depression (STD) and post-hypoxia frequency decline (PHFD)
Another time-dependent respiratory response during acute hypoxia is STD [61]. Although similar to HVD, STD was tentatively classified as a distinct mechanism by Powell et al. [49]. The primary difference is that STD is expressed as a decline in burst frequency (vs. tidal volume in HVD) and is seen within seconds to minutes of the acute hypoxic response [49,61]. STD has been reported only in rats and little is known about the underlying mechanism. However, a variant of STD is observed following a single hypoxic episode. A short-term reduction in respiratory burst frequency is often observed in anesthetized adult and geriatric rats, regardless of sex [61,81–84]. This decline in respiratory burst frequency is known as PHFD [81] and occurs due to an increase in the interval between inspiratory bursts (i.e., increased expiratory time) with little change in inspiratory burst duration. As such, the underlying mechanism may reflect potentiation of expiratory duration rather than inhibition of inspiratory drive per se [24]. Post-hypoxia frequency decline may not require carotid chemoafferent input since it is evoked by hypoxia in carotid denervated, awake rats [85]. However, persistent hypocapnia following hypoxia was not ruled out as a cause of PHFD in these experiments [85]. PHFD is abolished following lesions in the ventrolateral pons (A5 region; [81]). Moreover, A5 pontine electrical stimulation decreases inspiratory burst frequency in anesthetized rats [24]. Within the ventrolateral pons, noradrenergic neurons with respiratory related activity have been identified [86]. Taken together, the works of Guyenet et al. [86] and Dick and Coles [24] suggest that pontine adrenergic mechanisms are necessary for PHFD. Systemic a-2 adrenergic
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receptor antagonism attenuates PHFD in anesthetized rats [82], and PHFD is mimicked by systemic application of a-2 adrenergic agonists [87]. In contrast, intracerebroventricular injection of a different a-2 antagonist does not alter PHFD in rats [88]. Discrepancies between these reports may reflect different populations of laboratory rats (i.e., genetic differences; [89,90]), differences in experimental protocol, or differences in drug delivery, efficacy or specificity. Serotonergic and glutamatergic mechanisms may also influence PHFD. Pre-treatment with a 5HT2 receptor antagonist (ketanserin) accentuates PHFD in anesthetized rats [91]). The 5HT1 receptor agonist 5-carboxamidotryptamine also increased PHFD, an effect most likely attributable to diminished raphe neuron activity following activation of autoinhibitory serotonin type I receptors [91]. Excitatory glutamatergic transmission also influences PHFD because it is prevented by brainstem application of NMDA receptor antagonists [92]. Finally, PHFD in rats is subject to a degree of metaplasticity since prior hypoxic episodes diminish its expression following subsequent hypoxic exposures [82,93]. In summary, PHFD may result from complex interactions between different neuromodulatory systems (e.g., serotonergic, adrenergic, glutamatergic). The ventrolateral pons may be a critical neuroanatomical location for PHFD [24]. Brief Intermittent Hypoxia
When hypoxia is experienced in an intermittent or episodic pattern, unique forms of plasticity are revealed [4,19,94]. Following brief but continuous hypoxic exposures, respiratory motor output (electroneurograms or electromyograms) or ventilation return to pre-hypoxic baseline within minutes [37,94]. In contrast, brief periods of episodic hypoxia elicit a longlasting (minutes to hours) enhancement of inspiratory motor output known as respiratory long-term facilitation (LTF, Figure 6.2; [49,61]). LTF is usually expressed as increased inspiratory motor output or tidal volume in anesthetized animals [4,95], and as an increase in breathing frequency in awake animals [58,59,96,97]. Long-term facilitation is an excellent example of pattern sensitivity in respiratory plasticity since comparable periods of continuous hypoxic stimulation do not produce LTF [38,98,99]. The magnitude and pattern (augmenting vs. decrementing; tidal volume vs. frequency) of LTF are variable between preparations and laboratories. However, the fundamental observation is consistent: respiratory motor output (e.g., electroneurograms, electromyograms, tidal volume or frequency) remains elevated for a sustained period following episodic chemoafferent stimulation [4,95]. LTF has been demonstrated in sleeping humans [100–102], awake ducks [56], goats [96], dogs [103], rats [97,104] and mice [58], and anesthetized cats [52]
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Figure 6.2 Phrenic long-term facilitation (LTF). A. Representative tracing of phrenic LTF in an anesthetized, paralyzed, vagotomized, and ventilated rat. A typical experiment consists of three, 5-min bouts of isocapnic hypoxia followed by 1 h of isocapnic hyperoxia (lower tracing). Amplitude of the integrated phrenic activity (upper trace) increases during each hypoxic exposure, but returns to near baseline (dotted line) upon return to hyperoxia. Following the third hypoxic exposure, phrenic activity gradually increases; this increase in phrenic motor output is LTF. B. Working model for phrenic LTF. We propose that phrenic LTF results (at least in part) from potentiation of excitatory glutamatergic inputs to phrenic motoneurons and therefore represents a spinal mechanism (Data from Refs. 1, 4, and 95). In this model, intermittent hypoxia causes intermittent release of serotonin, thereby activating serotonin type 2 (5-HT2) receptors on phrenic motoneurons. Activation of metabotropic 5-HT2 receptors initiates intracellular signaling cascades that lead to increased translation of new proteins, such as BDNF. These proteins may act on the phrenic motoneuron, or on excitatory glutamatergic bulbospinal neurons, thus enhancing glutamate-dependent currents during inspiration. Similar mechanisms likely contribute to LTF in other respiratory motor outputs (e.g., hypoglossal, intercostal).
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and rats [4,95]. It should be noted that certain preparations or protocols have failed to yield LTF [104–106]. The reasons underlying this apparent contradiction are not clear in all cases, but LTF is susceptible to modulation and metaplasticity. Accordingly, the presence (or absence) of certain factors could eliminate (or enhance) LTF expression. For example, the magnitude of LTF is altered by the severity, duration, and number of hypoxic episodes [104,107] as well as the magnitude of the acute hypoxic response [95,108]. LTF is also influenced by age, sex, sex hormones, the estrus cycle [109,110], and genetics [90,111]. The probability of LTF also decreases as baseline respiratory motor output increases [95]. That is, if the difference in motor activity between baseline and maximum (e.g., maximal hypercapnic ventilatory response) conditions is (relatively) small, the capacity to exhibit plasticity may also be small (i.e., minimal LTF). Therefore, conditions which increase respiratory drive at baseline (e.g., mild hypercapnia; respiratory tract infections) may limit or prevent LTF. Our laboratory has observed on several occasions that respiratory viral or bacterial infections in our rat colony substantially reduce the probability of evoking phrenic LTF (Mitchell, G.S. and colleagues, unpublished observations). However, the latter observation remains anecdotal and awaits systematic investigation. Long-term facilitation is more robust in anesthetized preparations in which many confounding variables (e.g., PaCO2) can be rigorously controlled [95]. Vagal influences, which are often absent in reduced preparations, are inhibitory to or obscure LTF [112]. Nevertheless, LTF can be evoked in awake, vagally intact animals [59,96,97,104]. LTF has not been shown in awake humans [106], but can be activated during sleep [100,102]. In particular, sleeping humans who demonstrate significant inspiratory flow limitation develop substantial LTF following hypoxic episodes. LTF of upper airway muscle activity (e.g., genioglossus) has been postulated to stabilize breathing in patients with sleep-disordered breathing [49,102], consistent with recent reports of upper airway muscle LTF in humans [101,102,113]. Pharmacologic studies indicate that LTF is serotonin (5-HT) dependent [91,114–116]. Further, serotonin 5-HT2 receptor activation is necessary during (but not following) bouts of hypoxia [117]. Thus, 5-HT2 receptor activation during hypoxia initiates events that lead to, and maintain LTF. This observation reconciles potential inhibitory effects of serotonin [118] with serotonin-dependent facilitation. Inhibitory effects (possibly via pre-synaptic 5-HT1 receptors [119], may be restricted to the period of serotonin exposure, whereas the excitatory effects initiated by 5-HT2 receptor activation persist following receptor activation [117]. Brain-derived neurotrophic factor (BDNF) plays a critical role in LTF in anesthetized rats [4]. BDNF is necessary and sufficient for some forms of synaptic plasticity, and BDNF increases serotonin levels and turnover (reviewed in Ref. [4]). Episodic hypoxia increases BDNF in the
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ventral cervical spinal cord through a serotonin-dependent, protein synthesis-dependent mechanism [120]. Further, BDNF seems to be necessary and sufficient for phrenic LTF because pharmacological blockade of its receptor (TrkB) prevents LTF and spinal application of BDNF mimics LTF [120a]. Long-term facilitation may require NO [19,59], a molecule that also contributes to breathing stability during hypoxia [69,121]. Awake mice show LTF following 3 episodes of hypoxia (7% O2), however, LTF is not present following pharmacological block of NOS-1 or in NOS-1 knockout mice [59]. Kline et al. [59] suggest that nitric oxide may permit LTF by modulating 5-HT release from raphe neurons or by prolonging the actions of 5-HT on respiratory neurons. Although NO may increase with 5-HT2 receptor activation [122], nitric oxide may also inhibit 5-HT2 receptor function [123]. Thus, the role of nitric oxide in LTF will require further investigation. Where does the mechanism of LTF reside? The seminal experiments by Millhorn et al. [114,124] suggest that LTF can be accounted for by a central neural mechanism because it was evoked by CSN electrical stimulation. However, LTF may not require carotid chemoafferent inputs since a form of LTF can be evoked by intermittent hypoxia in carotiddenervated rats, although the amplitude is significantly reduced [108]. Morris, Lindsey and colleagues [10,45,46,125,126] suggest that LTF may be a consequence of pre- and/or post-synaptic changes in effective connectivity between inspiratory driver and premotoneurons in the medulla. Their data show that pontomedullary neural networks generate spatiotemporal patterns of synchrony, and enhanced synchrony may contribute to LTF [10]. However, responses of multiple respiratory neurons have been recorded only during the initial minutes of LTF [10]. The putative contribution of altered functional connectivity in the medulla to sustained LTF is unknown. While LTF can be evoked in different respiratory motor outputs (e.g., hypoglossal, phrenic, intercostals), a growing body of evidence suggests that in the case of phrenic activity, serotonin may act via spinal mechanisms to initiate phrenic LTF. First, hypoxia elicits serotonin release in the phrenic motor nucleus (detected by carbon fiber electrodes; [23]) and serotonergic terminals are present in the region of the phrenic motor nucleus (for review see Ref. [118]). Second, spinal application of methysergide (a serotonin receptor antagonist) or protein synthesis inhibitors blocks phrenic but not hypoglossal LTF [120], indicating a spinal site of action. Third, short latency phrenic potentials evoked by spinal stimulation at C2 are augmented following LTF [127]. Finally, a pre-conditioning lesion (cervical dorsal rhizotomy—CDR) that increases serotonin terminal density near identified phrenic motoneurons also enhances phrenic LTF in anesthetized rats [128]. Collectively, these observations support the hypothesis that phrenic LTF results (in large part) from spinal cord mechanisms. However, this
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hypothesis does not preclude additional effects at supra-spinal or peripheral locations [4,10,19,114,124]. We recently proposed working network and cellular/synaptic models of phrenic LTF (Figure 6.2; [4,5,116,118]. First, caudal raphe neurons projecting to the brainstem and spinal cord are activated during hypoxia [129,130]. Serotonin release from raphe neurons [23] during (but not following) hypoxia [117] activates post-synaptic 5-HT2A receptors on phrenic motoneurons [131], leading to kinase activation and new protein synthesis (translation from existing mRNA). These new proteins enable LTF by strengthening descending excitatory bulbospinal inputs to phrenic motoneurons [4,120]. Since bulbospinal respiratory drive in adult rats involves both NMDA and non-NMDA receptors [132,133], we postulate that newly synthesized BDNF phosphorylates glutamate receptors in the spinal cord [134]. By phosphorylating glutamate receptors, increased currents or insertion of additional glutamate receptors into the postsynaptic membrane may result [135]. Potentiated glutamate receptor currents would amplify descending respiratory drive, increasing phrenic output for the same pre-synaptic glutamate release (i.e., LTF). BDNF could also increase pre-synaptic glutamate release [136]. Sustained Hypoxia (41 h)
Detailed reviews of this topic can be found in Dempsey and Forster [137], Bisgard and Neubauer [37], Bisgard and Forster [75], Bisgard [18] and Powell et al. [49,138]. Hypoxic exposures greater than 1 h lead to at least two time-dependent ventilatory responses. The first response is a progressive increase in ventilation that reaches a plateau within hours to days, depending on the species (ventilatory acclimatization to hypoxia, VAH; [49]). The new, steady-state ventilation often exceeds the acute hypoxic ventilatory response prior to VAH [37]. The second mechanism is revealed as a decrease in hypoxic chemosensitivity in long-term high altitude residents, and is not considered further in this section (for review see Ref. [139] Chapter 11 by Powell and Bickler in this volume). Ventilatory acclimatization to hypoxia is characterized by a progressive decrease in PaCO2 due to the increase in minute ventilation during hypoxia. The length of time required for complete VAH is variable among species, ranging from hours to days [37,75], potentially reflecting distinct mechanisms or different time courses of the same general mechanism. Bisgard [18] provides an overview of peripheral chemoreceptor function during and following sustained hypoxia. Goats experiencing carotid body hypoxia with systemic (including CNS) normoxia exhibit VAH [140]. VAH is unique to hypoxia in goats, as perfusion of isolated carotid bodies with hypercapnic blood did not evoke this response [141]. Further, progressive increases in carotid chemoafferent activity are observed
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during VAH in anesthetized goats [142] and cats [143]. Thus, carotid body mechanisms contribute to (or are solely responsible for) VAH in certain preparations [18]. What mechanisms are active within the carotid body? Carotid body neuropeptides, such as endothelin, change in a manner consistent with increased carotid body sensitivity during sustained hypoxia [144–146]. In contrast, carotid body dopaminergic and noradrenergic mechanisms may be upregulated during sustained hypoxia, changes that would seem to favor a reduction in carotid chemosensitivity [146a]. Thus, the increase in carotid body hypoxic sensitivity may reflect a complex balance between excitatory and inhibitory neuromodulators [18]. Powell et al. [138] review the evidence for CNS mechanisms of VAH. They propose that a given chemoafferent input to the CNS produces a greater hypoxic ventilatory response following sustained hypoxia (e.g., days). This suggestion is supported by data from rats showing that integrated hypoxic phrenic responses to chemoafferent nerve activation are enhanced following days to weeks of hypoxia [138,147]. Neurochemical mechanisms of this plasticity are unknown. Serotonin, which enables respiratory LTF following intermittent hypoxia, probably does not play a critical role in VAH [118]. Huey and Powell [148,149] investigated the role of dopamine in VAH. After an extensive series of experiments on rats, they concluded that alterations in dopaminergic function do not explain increased CNS gain of the hypoxic ventilatory response after sustained hypoxia. In summary, sustained hypoxia induces plasticity, manifest as an increase in the acute hypoxic ventilatory response. This process may be initially dominated by peripheral chemoreceptor sensitization [37], followed by progressively increasing contributions from the central neural integration of carotid chemoafferent neurons [147,150,151]. Chronic Intermittent Hypoxia (CIH)
Chronic intermittent hypoxia is a more common experience than sustained hypoxia [19]. The physiological effects of CIH have been studied since the 1930s [152]. Scientists in the former Soviet Union initiated these studies in an attempt to better prepare Soviet pilots for flights in open cockpits up to 6000 m. Serebrovskaya [152] provides an excellent summary of this early Soviet research. CIH reportedly has been used as a treatment for disorders ranging from depression to radiation sickness, although the mechanistic bases for these reported effects are unclear [152]. CIH has also been reported to enhance athletic performance [153]. However, there may be shortcomings that limit or constrain the potential of CIH as a therapeutic tool. For example, certain CIH protocols elicit pathophysiology such as systemic hypertension [154], altered sympathetic chemoreflexes [155] and hippocampal apoptosis [156,157].
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Plasticity induced by CIH may be determined by the pattern and duration of hypoxic exposures [4,19]. Three patterns of CIH will be discussed: 1.
Repeated hypoxic episodes lasting 5 1 min separated by periods of normoxia ranging from seconds to minutes. This pattern is similar to what is experienced by patients with central or OSAs. As a result, several laboratories have developed CIH protocols within these general guidelines [19,154]. 2. Hypoxic episodes lasting minutes separated by similar durations of normoxia. This type of CIH protocol has been used extensively in studies of respiratory neuroplasticity [4,158]. 3. Long-lasting (hours to days) periods of hypoxia separated by similar periods of normoxia (e.g., repeated, prolonged ascent to altitude).
CIH with short-duration hypoxia (51 min)
This pattern of CIH exposures is produced in the laboratory in an attempt to mimic the clinical condition of sleep apnea. However, apneic episodes in OSA patients are accompanied by mild hypercapnia as well, in effect producing an intermittent asphyxia (i.e., hypoxia and hypercapnia). Accordingly, it is important to ascertain differences between poikilocapnic vs. isocapnic or hypercapnic hypoxia, similar to OSA patients. Interestingly, the magnitude of post-apnea ventilation is highly correlated with the CO2 load during the preceding apneic episode [159]. This observation led to the suggestion that a CO2-dependent facilitation of respiratory motor output may occur transiently after apneic episodes [159]. Currently, the impact of OSA on respiratory control is controversial [19]. Some investigators report that ventilatory chemoresponses are enhanced in OSA patients, possibly contributing to ventilatory instability [160], whereas others report either no change [161,162] or a reduction in ventilatory chemosensitivity in OSA patients [163]. Peng et al. [107] measured hypoxic ventilatory responses following 10 days of CIH in rats (5% O2 for 15 sec, alternating with 21% O2 for 5 min, 8 h/day, 10 days). However, in this protocol, the 15-sec period of 5% O2 is followed by 4150 seconds of transient hypoxia. Rats were subsequently anesthetized and phrenic nerve activity was monitored during hypoxia. Rats treated with CIH had an almost 50% greater phrenic motor output during hypoxia than controls. This increase in the hypoxic phrenic response was mirrored by increases in carotid body chemosensitivity. Thus, the augmented hypoxic response could be accounted for, at least in part, by peripheral (carotid body) mechanisms. These data do not rule out central neural mechanisms [93]. Peng et al. [107] reported additional plasticity evoked by CIH. A robust (4200%) LTF of carotid chemoafferent activity
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could be evoked by episodic hypoxia (15 sec 5% O2, 5-min normoxia, 5-min intervals) in CIH-treated, but not control rats [107]. No obvious changes in carotid body morphology were found. Similar CIH protocols cause systemic hypertension in rats, although respiratory responses were not evaluated in these studies (reviewed in Ref. [154]). CIH with moderate-duration hypoxia (1 to 20 min)
Our laboratory developed a model of CIH in which rats experience one week of nocturnal intermittent hypoxia (11–12% O2/air at 5-min intervals, 12 h/night; [93]). The short-term and long-term phrenic responses to additional hypoxic episodes were examined in anesthetized, paralyzed, vagotomized, and artificially ventilated rats. CIH augments the short-term hypoxic phrenic response (during hypoxia), abolishes PHFD and enhances phrenic LTF by more than 100% [93]. Since these effects are reversed by methysergide, serotonin receptor activation is necessary to maintain these forms of plasticity [93]. However, ketanersin, a more selective 5-HT2 receptor antagonist, was less effective, suggesting an involvement of non5-HT2 serotonin receptors [93]. Since integrated phrenic responses to CSN stimulation are amplified following CIH [93], CIH-induced plasticity results, at least in part, from a central neural mechanism. This pattern of CIH may have therapeutic value, as it can restore blunted hypoxic phrenic responses following neonatal hyperoxia [164] and enhances existing, but ineffective, synaptic pathways below spinal cord injury [30]. Most studies of human responses to CIH have employed relatively short bouts of CIH (e.g., 1 h) repeated daily. The following CIH protocols have all been reported to enhance subsequent hypoxic ventilatory responses: 3 6 min hypoxia (end-tidal PO2 ¼ 35–50 mm Hg) separated by 4 min normoxia, 14 consecutive days [165]; 3–4 7 min progressive hypoxia (end-tidal PO2 reaching 35–40 mm Hg), 14 consecutive days [166]; 20 min isocapnic hypoxia per day, 14 consecutive days [167]. While these studies did not address the underlying mechanisms, they document that the hypoxic ventilatory response is subject to plasticity following CIH in humans. CIH with long-duration hypoxia (hours – days)
Several investigations have exposed humans to hypoxia for 1–2 h per day for approximately one to two weeks [152,168–170]. While these studies have protocol differences, they all report that the hypoxic ventilatory response was significantly enhanced following CIH. Another study followed Chilean miners commuting to work from sea level to 4500 m on a weekly basis [171]. A type of ventilatory acclimatization, characterized by an increased ventilatory response to hypoxia, began to emerge after 12 months of this CIH pattern. Elite athletes using the live high, train low approach [153] are another population exposed to CIH. There is some evidence that spending nights in simulated altitude environments and days at sea level (where the
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acute effects of altitude will not impair training) can improve athletic performance, presumably through adaptations in oxygen transport (e.g., increased red blood cell number). This pattern of CIH may also induce plasticity in respiratory control. For example, this pattern of CIH enhances acute hypoxic responses in humans [172]. To our knowledge, LTF has not been assessed following CIH in humans. In summary, neuroplasticity following CIH of various durations and patterns is manifest as an increase in the acute hypoxic response; most investigations suggest that the hypercapnic ventilatory response is not affected. CIH also induces a form of metaplasticity by enhancing LTF. The mechanisms underlying these changes may not be consistent across CIH paradigms. For example, repeated, short, intense hypoxic exposures may preferentially evoke peripheral (carotid body) plasticity while chronic exposure to longer, more moderate hypoxic bouts may induce central neuroplasticity. Confirmation of this hypothesis and elucidation of the detailed mechanisms underlying the effects of CIH will require further investigation. B. Hyperoxia
In mammals, hyperoxia (i.e., FiO2 4 0.21) is experienced only in experimental and clinical therapeutic conditions. There are no naturally occurring causes of hyperoxia in humans. Brief hyperoxia has been shown to induce plasticity in respiratory control [173,174]. A brief bout of hyperoxia (10 min, 100% O2) can enhance hypoxic ventilatory responses assessed shortly thereafter [173]. These authors speculated that the increased hypoxic ventilatory response reflected enhanced glutamate release in the nucleus of the solitary tract following hyperoxia. Gozal [174] extended these observations by demonstrating that a 10-min pre-exposure to hyperoxia (100% O2) caused a nitric oxide-dependent augmentation of the hypoxic ventilatory response in rats. Gozal speculated that activation of NMDA receptors led to NOS activation and potentiation of the hypoxic response. Sustained hyperoxia (100% O2; 12–60 h) blunts subsequent hypoxic responses in rats and cats [175–177] by reducing carotid but not aortic chemosensitivity [177,178]. The carotid body undergoes striking morphological changes during prolonged hyperoxia including type I cell necrosis [179,180]. Ren et al. [181] reported that 8 h of hyperoxia (end-tidal O2 ¼ 300 mm Hg) blunts the hypoxic ventilatory response in humans. In contrast, prolonged hyperoxia, either sustained or intermittent, does not blunt the hypoxic ventilatory response in humans [182]. Many studies of respiratory LTF have used anesthetized rats or cats with hyperoxic baseline conditions [83,84,90,94,116,117]. In these studies, the animals were ventilated with hyperoxic gas mixtures (FiO2 0.5) to prolong viability of the experimental preparation. One consequence of using
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a background of hyperoxia in episodic hypoxia studies is that, in addition to hypoxic episodes, animals are exposed to rapid rises in oxygen to hyperoxic levels at the end of each hypoxic bout. This led to an investigation of the effects of repeated, brief exposures to hyperoxia on resting respiratory motor output [108]. Although there was no evidence for time-dependent changes in phrenic activity following episodic hyperoxia (FiO2 ¼ 1.0), this finding does not preclude the possibility that episodic hyperoxia and/or exposure to reactive oxygen species (ROS) initiates other forms of respiratory plasticity. C. Hypercapnia
Hypercapnia results when alveolar ventilation is low relative to metabolic rate, such as following CNS injury (e.g., spinal cord injury), during central or OSAs, or chronic pulmonary diseases that exhibit CO2 retention (e.g., chronic obstructive pulmonary disease). Increases in inspired CO2 represent a potential cause of hypercapnia in closed environments (e.g., submarines, spacecraft) and are experienced by many burrowing species [183,184]. Although CO2 is a powerful respiratory stimulus and a primary determinant of respiratory motor output [185], it may also elicit inhibitory mechanisms of respiratory neuroplasticity as described below. Acute Sustained Hypercapnia
The initial ventilatory response to hypercapnia is an increase in tidal volume or its neural equivalent (phrenic neurogram or diaphragm EMG) with a lesser increase in burst frequency. Tidal volume generally reaches a plateau within a few minutes, and does not show much, if any, roll-off within this time frame. However, following a 25-min bout of hypercapnia, phrenic motor output was depressed for up to an hour in anesthetized rats [38]. This long-term depression (LTD) was evident as depressed phrenic burst amplitude for up to 60 min following hypercapnia. Phrenic burst frequency exhibited a transient (15-min) depression following hypercapnia, but then returned to baseline values. The mechanisms underlying LTD following sustained hypercapnia are pattern-sensitive since intermittent hypercapnia (of comparable cumulative duration) did not produce significant LTD in the same rat substrain [38]. Acute Intermittent Hypercapnia
Dong et al. [186] exposed anesthetized cats to intermittent 5% CO2 challenges. Facilitation of inspiratory output was seen during successive bouts of hypercapnia as inspiratory tidal volume and diaphragm electromyogram (EMG) activity increased despite concurrent decreases in frequency. Optical reflectance of the ventrolateral medullary surface was
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measured to provide an index of respiratory neural activity. Successive hypercapnic challenges increased the optical reflectance, suggesting that neural activity was enhanced by successive hypercapnic episodes. Unfortunately, long-term changes in respiratory motor output following episodic hypercapnia were not assessed [186]. Morris et al. [46] recorded phrenic motor output following episodic intracarotid injection of CO2-saturated saline. This intermittent hypercapnia protocol induced persistent facilitation of respiratory motor output similar to hypoxia-induced LTF (see above). However, intracarotid injections directly activate only carotid chemoreceptors with little, if any, direct effects on the central nervous system. Accordingly, these data do not reflect the effects of systemic or CNS hypercapnia on respiratory motor output. Episodic systemic hypercapnia induces LTD of respiratory motor output in anesthetized rats [187]. In this study, LTD was prevented or attenuated if rats were pretreated with a2-adrenergic receptor antagonists. Subsequently, Baker et al. [38] found that a similar pattern of hypercapnia caused a (non-significant) trend for LTD in anesthetized rats of a different substrain of Sprague–Dawley rats, suggesting possible genetic influences on LTD. Gozal and colleagues examined the ventilatory response to intermittent hypercapnia in awake humans [188,189]. Over the course of six, 2-min hypercapnic episodes (5% CO2) a significant shift in the pattern of breathing (increased tidal volume, decreased breathing frequency) with no change in the overall minute ventilation occurred. This effect was abolished if the interval between hypercapnic episodes was extended from 5 to 15 min, indicating that the underlying mechanism was sensitive to interval time. A similar change in breathing pattern during episodic hypercapnia occurs in healthy children, but not in pediatric OSA patients [189]. Gozal et al. speculate that the altered ventilatory response in pediatric OSA patients reflects plasticity induced by repeated asphyxic events during OSA. Thus, repeated episodes of hypercapnia (i.e., during apneic episodes) may change the subsequent pattern of ventilatory response to hypercapnia. Chronic Sustained Hypercapnia
The ventilatory response to chronic hypercapnia was investigated by exposing humans to 1.5% CO2 for 42 days [190]. Minute ventilation and alveolar CO2 were increased throughout the entire CO2 exposure although there was little change in these variables after the first few days. Alveolar CO2 and minute ventilation remained significantly elevated for 10 days upon return to normoxia. Dempsey and Forster [137] reviewed the existing literature on sustained hypercapnia in mammals. They concluded that the ventilatory response to chronic hypercapnia is biphasic [137,191]. After the acute onset of hypercapnia, ventilation remains elevated, but declines
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somewhat during subsequent days to weeks of hypercapnia. Recent studies in rats confirmed this conclusion [192] and indicate that ventilation during chronic hypercapnia follows the normal circadian rhythm [193]. Although arterial CO2 remains elevated for the duration of hypercapnia, ventilation, plasma and CSF pH can all return toward normal [137]. Accordingly, the mechanism for respiratory stimulation during chronic CO2 exposure remains controversial. During chronic hypercapnia, ventilatory sensitivity to CO2 decreases in mammals and birds [137,194], although exceptions have been reported [195]. Following chronic hypercapnia, ventilation and arterial CO2 can remain elevated for several weeks in humans upon return to normoxia [190]. However, CO2 chemoresponses return to control values within three to four weeks after return to normoxia [190]. Chronic Intermittent Hypercapnia
To our knowledge, there are no studies directly addressing the impact of chronic intermittent hypercapnia on respiratory plasticity. III.
Developmental Plasticity and the Control of Breathing
The expression of plasticity may depend on the age and/or developmental stage during which the environmental or experimental stimulus is presented. Developmental plasticity refers to plasticity that is specific to periods of development [1,6]. Although plasticity is not unique to development, immature animals may exhibit greater capacity for plasticity, including forms of plasticity that cannot be elicited in the mature animal or that persist for longer periods (e.g., weeks to years). This developmental specificity suggests the existence of critical periods (also known as developmental windows, Figure 6.3), when the phenotype is particularly sensitive to prevailing external or internal conditions. Critical periods may vary in their timing, duration and underlying mechanisms [196]. Understanding developmental plasticity is critical to appreciate normal development of respiratory control mechanisms, various respiratory disorders (e.g., sudden infant death), as well as phenotypes associated with altitude or burrowing life-styles. In the following sections, we briefly review developmental plasticity in mammalian respiratory control induced by altered respiratory gases, altered metabolic rates, neural injury (i.e., chemoafferent denervation), and stress. Further, we consider how this plasticity may relate to pathophysiology. Developmental plasticity in respiratory control is not limited to mammals and has been demonstrated in a variety of vertebrate and invertebrate species, emphasizing that this topic has evolutionary as well as biomedical significance.
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A. Time 250 200 (∆bl, %bl)
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l % 0% % % % tro 0% 60 60 6 60 60 ts) 6 2 4 3 1 4 a 4 – r ks ek ek ek 1– s 1 atric ee We We We k eri ks e W e e g e W ( W n Co
Figure 6.3 A. During development, there are critical periods (also known as developmental windows) during which respiratory phenotypes are sensitive to environmental conditions. B. The critical period for hyperoxia-induced developmental plasticity in rats extends into the second postnatal week. The figure shows phrenic nerve responses to isocapnic hypoxia (PaO2 40 mm Hg) in anesthetized rats exposed to 1–4 weeks of developmental hyperoxia (60% O2); control animals were raised in normoxia. Adult (3–5 month) and geriatric rats (14–15 month) exposed to 4 weeks of hyperoxia from birth have substantially blunted hypoxic phrenic responses. Adult rats exposed to developmental hyperoxia for either the first or second postnatal week (but not the third or fourth week) also have blunted hypoxic responses. Accordingly, the critical period for this plasticity includes the first two post-natal weeks. It is not known whether the critical period extends prenatally as well (figure compiled from previously published data). (Data from Refs. 251, 252.)
A. Prenatal Hypoxia
Exposing pregnant dams to the equivalent of 12% O2 throughout gestation causes persistent hyperventilation in the newborn rat [197]. These effects are similar to VAH in adult rats [198], although somewhat greater effects on metabolism are observed. Resting ventilation and the ventilation to metabolism ratio remains elevated at 1 and 3 (but not 9) weeks of age following prenatal hypoxia (10% O2; [199]). Thus prenatal hypoxia induces persistent hyperventilation of longer duration than VAH in adults [137].
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Prenatal hypoxia also has lasting effects on hypoxic ventilatory and metabolic responses [199], generally enhancing ventilation and abolishing metabolic changes during hypoxia at 1 and 3 weeks of age. These effects are transient, however, and the hypoxic ventilatory response is attenuated relative to normal by 9 weeks of age [199]. It is not known whether altered normoxic and hypoxic ventilation following prenatal hypoxia reflect plasticity in the control of breathing per se since blood gases were not measured in these studies; hypoxia-treated rats could have experienced more severe hypoxia and/or hypercapnia due to impairments in gas exchange. Effects of prenatal hypoxia could also result from changes in respiratory mechanics [200]. However, prenatal hypoxia alters the postnatal expression of dopamine, noradrenaline, and related enzymes in the carotid body and brainstem respiratory centers [199,201], suggesting neuroplasticity. Further, carotid body glomus cells isolated from neonatal rats following prenatal hypoxia (second day of gestation through postnatal days 5–8) exhibit membrane current properties consistent with increased excitability [202,203]. While it is unknown if these carotid body effects persist beyond the first few days post-hypoxia, these data suggest a mechanism for enhanced hypoxic responses following prenatal hypoxia. Most studies of prenatal hypoxia produce fetal hypoxemia by exposing the mother to environmental hypoxia (i.e., low inspired PO2), as may occur at altitude. However, plasticity in the control of breathing may result from other sources of prenatal hypoxia. Intrauterine growth restriction, such as that caused by chronic placental insufficiency or experimental manipulation (e.g., ligation of the uterine artery, maternal anemia), may result in fetal hypoxemia. Intrauterine growth restriction attenuates the postnatal hypoxic ventilatory response in young sheep during wakefulness [204,205], but not during sleep [206,207], and may increase the ventilatory response to progressive asphyxia (i.e., hypercapnic hypoxia) in newborn guinea pigs [208]; none of these studies found a change in steady-state hypercapnic ventilatory responses. Prenatal exposure to carbon monoxide (CO) produces tissue hypoxemia by reducing O2 content of fetal blood. In guinea pigs, repeated exposure to prenatal CO (10 h/day) for the last 6 weeks of gestation increases hypercapnic ventilatory responses in 4–5 day old pups, despite normal resting ventilation [209]. Additional studies are needed to verify that fetal hypoxemia is the common stimulus in these examples, but collectively, these data suggest that prenatal hypoxia can have lasting effects on control of breathing. B. Sustained Neonatal Hypoxia
Neonatal hypoxia induces short- and long-term respiratory plasticity distinct from adult plasticity. For example, rats raised in hypoxia (10% O2) for the first postnatal week hyperventilate when transferred to room
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air ([210]; see also [211]). Hyperventilation is persistent, lasting 6–7 weeks after return to normoxia despite normal PaO2 [212,213]. A similar hypoxic exposure in post-puberty rats has no enduring effects, indicating that the critical period for this plasticity does not extend into adulthood [212,213]. Whereas prenatal and adult exposures to chronic hypoxia generally enhance acute hypoxic ventilatory responses, hypoxia during early postnatal development appears to blunt hypoxic responses. Indeed, chronic neonatal hypoxemia may explain blunted hypoxic responses in humans born with cyanotic heart disease [214–216]. Humans born in hypobaric hypoxia also exhibit reduced hypoxic ventilatory responses as adults, even after moving to sea level [139,217]. This effect can only be partially attributed to genetic mechanisms [139]. For example, high-altitude residents may be born with normal hypoxic responses and gradually acquire blunted responses during postnatal development [139,218–221]; similar changes are not observed, or occur more slowly and to a lesser extent, in sea level natives after moving to altitude as adults [218,219,221]. How long hypoxic ventilatory responses remain blunted once normoxia is restored is unclear [139,214–218,222], though any recovery appears to be gradual. Hypercapnic ventilatory responses are generally reported to be unaltered by neonatal hypoxia in humans [214,218,219,223], although other studies have found both reduced [224] and enhanced [225] ventilatory responses to CO2 in high-altitude natives. Similar effects of neonatal hypoxia on hypoxic ventilatory responses have been observed for non-human mammals, confirming long-lasting developmental plasticity. In young rats, cats and sheep born and raised in hypoxia, ventilatory responses to acute hypoxia are abolished or greatly reduced 0–24 h after return to normoxia [226–231]. In rats and sheep, hypoxic ventilatory responses remain blunted for weeks to months following 1–2 weeks of neonatal hypoxia despite normal blood gases and/or metabolic rates [213,229,232]. In rats, this long-lasting plasticity cannot be elicited after puberty [213] and is expressed only by males [232]. Hypercapnic ventilatory responses are generally normal following neonatal hypoxia [213,234,235], indicating that the effects of neonatal hypoxia are specific to the hypoxic response. The mechanism(s) for long-lasting attenuation of the hypoxic ventilatory response following neonatal hypoxia are unclear. Neonatal hypoxia (from birth) delays maturation of carotid body responses either to hypoxia [228,230,235,236] or to combined hypoxia and hypercapnia [237]. However, even if rats are born and raised in hypoxia, carotid body O2 chemosensitivity appears to recover spontaneously despite continued hypoxia [226] or following return to normoxia [235,236]. Moreover, phrenic nerve responses to isocapnic hypoxia are unaltered in adult male rats raised in hypoxia for postnatal week 1, despite having blunted hypoxic ventilatory
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responses [232]. Thus, changes in peripheral chemosensitivity, chemoafferent pathways or central neural integration of chemoafferent inputs cannot explain the long-lasting attenuation of hypoxic ventilatory responses following neonatal hypoxia; the impairment appears to be downstream of phrenic activity (Figure 6.1). Consistent with this conclusion, neonatal hypoxia alters respiratory system mechanics and lung morphology [212,238], and diaphragm function [31,239]. Thus, neonatal hypoxia may have short-term effects on carotid bodies whereas anatomical and/or neural changes in respiratory muscles or respiratory mechanics may be responsible for long-lasting plasticity. C. Intermittent Neonatal Hypoxia
Intermittent hypoxia is observed clinically, often in combination with mild hypercapnia due to periodic apneas or impaired gas exchange (e.g., OSA, apnea of prematurity, asphyxia during birth, bronchopulmonary dysplasia). In neonatal rats, pre-treatment with short periods of intermittent hypoxia reduces HVD during subsequent exposures [121,158]. Thus, intermittent hypoxia enhances hypoxic ventilatory responses in both neonatal and adult rats [93,158]. This facilitatory plasticity may diminish with longer periods of intermittent hypoxia in both neonates and adults [19,158]. In experimental studies with rats, both prenatal and neonatal CIH impairs anoxic gasping [240]. Moreover, 30 min of severe hypoxia (6% O2) twice daily for the first four postnatal days reduces the hypoxic ventilatory response one day later in rats, but these effects are no longer evident at two weeks of age [241]. Consistent with rapid recovery following neonatal intermittent hypoxia, CIH during development does not appear to alter the adult hypoxic ventilatory response [242]. However, while daily bouts of anoxia (100% inspired N2) had no detectable effect on the hypoxic ventilatory response at 7 days of age [243], a single 20-min anoxic episode in 3–4 day old neonatal rats enhanced the hypoxic ventilatory response measured at 25 days of age [244]; no effects on the hypoxic response were detected at 9 days of age. Thus, even a solitary severe hypoxic episode during the neonatal period can have lasting effects on the control of breathing that are only revealed later in life, emphasizing that multiple post-exposure time points may be necessary to adequately assess the expression of developmental plasticity. Several studies have considered the effects of intermittent hypoxia during development in piglets. Piglets (3–5 week old) exhibit blunted hypoxic ventilatory responses following five daily exposures (30 min/d) to poikilocapnic hypoxia [245], consistent with data in rats [241]. However, there is some evidence that rapid cycles of hypoxia within a single day (7 cycles, 3 min hypoxia/3 min air) also attenuate the hypoxic ventilatory response in this species at 2–3 week of age [246]. Since adults of other species
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exhibit enhanced hypoxic sensitivity following intermittent hypoxia [19,93,158], this manifestation of plasticity may be unique to development in piglets. However, until similar studies are completed in adult pigs, it remains possible that blunted hypoxic responses following intermittent hypoxia is a species-specific trait. A very different picture emerges when neonatal piglets (1–2 weeks old) are exposed repeatedly to hypercapnic hypoxia [247]. Acute hypoxic ventilatory responses, measured 24 h following the last of seven daily exposures to 10% O2 and 6% CO2, are enhanced in piglets despite normal blood gases; acute responses to hypercapnia or hypercapnic hypoxia are generally reduced. Moreover, the pattern of hypercapnic hypoxia during each daily episode (intermittent vs. continuous) appears to influence the resulting plasticity [247]. Thus, the effects of CIH during development on ventilatory control are complex, depending on the exposure pattern and duration, background PaCO2 and species studied. D. Hyperoxia
Exposure to 30–60% O2 for a week or more during early (primarily postnatal) life reduces hypoxic ventilatory responses in rats and cats [228,248,249,249a]. In kittens born and raised in 30% O2 for the first 12–13 days of life, acute hypoxic ventilatory responses are absent immediately following the hyperoxic exposure [229]; it is not known if these effects are persistent. In contrast, rats exposed to 30% O2 for the first 5 days of life exhibit sustained increases in ventilation during acute hypoxia (vs. the biphasic hypoxic response typical of neonatal rats), suggesting accelerated maturation of the hypoxic ventilatory response [250]. However, the carotid body response to hypoxia was abolished in rats after 5–10 weeks of hyperoxia from birth in the same study [250]. It is possible that 5 days of 30% O2 is not long enough to blunt the hypoxic ventilatory response, or that the exposure period did not encompass the critical period for this plasticity (see below). Our laboratory has demonstrated that 1–4 weeks of developmental hyperoxia (30–60% O2) causes long-lasting attenuation of the hypoxic ventilatory and/or phrenic nerve responses in rats [17,164,248,249,249a,251,252]. After controlling for blood gases and metabolic rate, hypoxic ventilatory responses are significantly reduced despite normal normoxic ventilation in awake, adult rats (43 months of age) exposed to 60% O2 for the first week [249a] or month [248] of life, primarily due to a smaller increase in respiratory frequency; hypercapnic ventilatory responses are unaffected by developmental hyperoxia [248]. Phrenic responses to isocapnic hypoxia are also reduced in anesthetized rats exposed to hyperoxia during development [17,164,249,249a,251,252]. Although the effects of one month of 60% O2 on hypoxic phrenic responses appear permanent [251], rats experiencing shorter (i.e., one week) hyperoxic
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exposures exhibit a degree of spontaneous recovery [249a]. However, hypoxic responsiveness can be restored (at least temporarily) to rats exposed to one month of 60% O2 during development by treatment with chronic sustained or intermittent hypoxia as adults [164]. It is not known whether this functional recovery reverses the effects of developmental hyperoxia or enhances phrenic responses by inducing normal, adult-type plasticity of the residual hypoxic response (see section above on Chronic intermittent hypoxia). Adult rats exposed to 60% O2 for one month exhibit no lasting changes in ventilatory [248] or phrenic [249] responses to hypoxia, indicating that the plasticity following developmental hyperoxia is specific to development. A recent study revealed that the critical period for hyperoxia-induced respiratory plasticity extends into the second postnatal week ([252]; Figure 6.3); one week of 60% O2 has no lasting effect on the hypoxic phrenic response of rats treated during the third or fourth postnatal week. However, since hyperoxic exposures begin a few days prior to birth in most of these studies [17,164,248,249,249a,251,252], there is some possibility of pre-natal effects of hyperoxia as well. The effects of prenatal hyperoxia have not been studied explicitly. Nevertheless, prenatal hyperoxia is not necessary for this form of developmental plasticity since hypoxic responses are reduced in adult rats that have been exposed to 60% O2 during the second postnatal week only [252]. Since phrenic responses to isocapnic hypoxia are blunted in rats following developmental hyperoxia, the effect of hyperoxia on the hypoxic ventilatory response cannot be (wholly) explained by changes in respiratory muscles or respiratory mechanics. Moreover, electrical stimulation of the CSN (i.e., bypassing hypoxic chemotransduction in the carotid body) produces equivalent increases in phrenic motor output in hyperoxia-treated rats and untreated controls [251,253], indicating that central integration of afferent information from peripheral chemoreceptors is not impaired. Several lines of evidence indicate that developmental hyperoxia induces plasticity at the carotid body chemoreceptors. For example, carotid body morphology is altered by 1–4 weeks of 30–60% O2 during development, with reduced carotid body volume (total volume and volume occupied by glomus cells), fewer carotid chemoafferent neurons and altered dopamine content [164,251,254,255]. Attempts to measure hypoxic sensitivity of carotid body chemoreceptors through single fiber recordings have been unsuccessful in adult rats exposed to developmental hyperoxia [17], possibly reflecting the reduced number or fragile condition of chemoafferent neurons. In the absence of single unit recordings, carotid body responses have been assessed from whole CSN preparations. These studies consistently demonstrate reduced carotid body responses to hypoxia, asphyxia and/or intravenous cyanide injection [17,250,251,255,256]. It is not clear from these whole nerve studies whether individual glomus cells are less sensitive to
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hypoxia or if hypoxic responses are reduced because there are fewer glomus cells and/or chemoafferent neurons. However, it was recently reported that carotid body dopamine release is reduced during hypoxia, but not in response to extracellular Kþ, in adult rats several months after developmental hyperoxia [255], suggesting decreased hypoxic sensitivity. Moreover, immediately after 1–3 weeks of hyperoxia from birth, single unit carotid body responses to hypoxia are depressed in rats [257] and kittens [228], as are the depolarization and intracellular calcium responses to hypoxia in carotid body glomus cells isolated from hyperoxic rats [258]. Thus, developmental hyperoxia may attenuate hypoxic responses through lasting effects on both the absolute numbers and O2 sensitivity of carotid chemoreceptor cells. Conversely, since phrenic responses to CSN stimulation in rats exposed to developmental hyperoxia are normal [251,253], the reduced number of axons from chemoafferent neurons [254] may not be a major contributor to impaired hypoxic ventilatory responses. The pathways by which developmental hyperoxia alters carotid body function are currently unknown. By raising PaO2 above the threshold for carotid body activity, hyperoxia may reduce or prevent depolarization of the carotid body chemosensory cells. This may in turn diminish the activitydependent synthesis and/or release of neurotransmitters and trophic factors in the carotid body chemosensory system during critical periods of development [259,260]. For example, carotid chemoafferent neurons (and perhaps glomus cells themselves) require the release of trophic factors from intact carotid bodies during the early postnatal period [261]. The synthesis and release of trophic factors may be activity dependent [260] and there is suggestive evidence that trophic factors such as BDNF are reduced in the carotid body following chronic hyperoxia (unpublished data cited in Ref. [254]). Thus, carotid chemoreceptor inactivity could result in inadequate trophic factor availability during developmental hyperoxia, thereby causing carotid body hypoplasia and degeneration or withdrawal of chemoafferent neurons [254]. Carotid chemoafferent neurons no longer require trophic support by the third postnatal week, perhaps earlier, [261], consistent with the critical period for hyperoxia-induced changes in carotid body morphology [254] and hypoxic phrenic responses [252]. In addition to reducing neural activity, hyperoxia could alter the expression of genes regulated by O2. Many genes directly related to carotid body function are regulated by hypoxia, including genes related to O2 sensing (e.g., ion channels), neurotransmitters and neurotransmitter release [262], and these genes may also be susceptible to hyperoxia [263]. Other O2-sensitive genes and gene products could have indirect effects on carotid body function, such as vascular endothelial growth factor (VEGF), which has been identified in the carotid body (Wang, Z.-Y. and Bisgard, G.E., personal communication). If hyperoxia downregulates VEGF production in the carotid body,
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as it does elsewhere in the body [264], developmental hyperoxia could lead to poor carotid body vascularization. Loss of vasculature could alter tissue O2 profiles, or could cause a loss of glomus tissue (and thus trophic support for developing chemoafferent neurons). Finally, the effects of hyperoxia on carotid body function could be mediated by ROS, either through altered gene expression or cellular toxicity [263,265–269]. The neural effects of developmental hyperoxia (60% O2) seem to be specific to the carotid body and related chemoafferent pathways, with no impairment of neurons in the nodose ganglion [254], the hypercapnic ventilatory response [248] or the central neural integration of chemoafferent inputs [251,253]. Thus, the effects of developmental hyperoxia on respiratory control are not due to widespread, non-specific cellular toxicity. However, given its high blood flow relative to metabolic rate, the carotid body may be more susceptible to hyperoxia than other tissues [175,178]. Accordingly, local ROS-mediated toxicity could directly or indirectly lead to the loss of glomus cells and/or chemoafferent neurons. Developmental hyperoxia is rare in non-experimental settings, and is probably limited to mechanical ventilation and/or supplemental oxygen therapies in infants. Attempts are made to maintain SaO2 (and therefore PaO2) within limits to minimize or prevent hyperoxic toxicity and conditions such as retinopathy of prematurity. However, periods of hyperoxia are likely to occur during oxygen therapy [270], and the lower limits of hyperoxic exposure required for hyperoxia-induced respiratory plasticity are poorly understood. In human infants, there is a relationship between oxygen therapy and blunted hypoxic ventilatory responses [271]. It is not known whether this relationship is causal, or if hypoxic chemosensitivity was reduced in these infants due to prior exposure to intermittent hypoxia associated with bronchopulmonary dysplasia or septicemia [271–274], but this relationship highlights the need to understand the effects of oxygen therapy on ventilatory control from a biomedical perspective. Relative hyperoxia at (premature) birth (i.e., the abrupt rise from fetal PaO2) may also induce hyperoxia-related plasticity in respiratory control, either as part of normal development or pathologically, but this has not been studied [6]. E.
Hypercapnia
Although many burrowing mammals and birds regularly experience high levels of inspired CO2 during development [183,184], environmental hypercapnia is rare in humans, particularly during development. However, hypercapnia may occur during some diseases, often in conjunction with hypoxia (e.g., chronic lung disease). Moreover, current strategies for neonatal respiratory therapy may include permissive hypercapnia, a strategy
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in which low tidal volumes are used during mechanical ventilation to minimize lung trauma while allowing PaCO2 to rise to 45–55 mm Hg or higher [275]; the effects of permissive hypercapnia on the development of respiratory control have not been studied. The few studies examining developmental hypercapnia and the control of breathing in mammals have reached opposite conclusions. Birchard and colleagues [276] studied male rats exposed to 6% CO2 from fertilization through the third postnatal week (i.e., throughout prenatal and postnatal development). When studied after 6 weeks in room air, blood gases, metabolic rates and ventilation were similar to control rats at three levels of inspired CO2 (0–5%), indicating no long-lasting effects on the control of breathing. In a similar set of experiments, Rezzonico and Mortola [277,278] studied male and female rats exposed to 7% CO2 for six days (beginning 24 h after birth). Two days after the return to room air, CO2-treated rats exhibited greater resting ventilation and reduced ventilatory responses to 10% inspired CO2 (expressed as percentage increase from baseline) [278]. When studied at 45–50 days of age (i.e., 6 weeks post-exposure), CO2-treated rats had reduced resting ventilation, normal metabolic rates, normal hypoxic ventilatory responses (10% O2), and reduced hypercapnic ventilatory responses (10% CO2) [277]. Thus, in contrast to the earlier study by Birchard and colleagues, this study found evidence for long-lasting plasticity in respiratory control following developmental hypercapnia. Several methodological differences could contribute to these divergent conclusions, including the timing, duration and severity of developmental hypercapnia and the levels of CO2 used to assess the hypercapnic ventilatory response. Alternatively, the differences may be explainable by the sex of the rats studied. In female Japanese quail (Coturnix japonica) treated with 2% CO2 throughout embryonic development, adult hypercapnic ventilatory responses are reduced despite similar blood gases, metabolic rates, and resting ventilation [279]; however, the hypercapnic ventilatory responses of male quail were unaffected by developmental CO2. Re-analysis of an earlier study [280] revealed a similar pattern for zebra finches (Taeniopygia guttata): only female finches exhibit blunted hypercapnic ventilatory responses following embryonic CO2 exposure [279]. Thus, sex may have profound effects on the expression of developmental plasticity and may contribute to the different conclusions reached in studies with rats. Indeed, Birchard and colleagues [276] studied only male rats, whereas Rezzonico and Mortola [277,278] report data for a combination of males and females (Mortola, J.P., personal communication). Acclimation to CO2 as adults produces only transient effects on the hypercapnic ventilatory response (reviewed above), suggesting that the longlasting effects (weeks to months) of developmental CO2 exposure in rats and birds are specific to development. No studies have attempted to define the critical period for this CO2-induced developmental plasticity, although data
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from birds suggest that both embryonic (i.e., prenatal) and nestling (i.e., postnatal) CO2 exposures produce qualitatively similar effects on the hypercapnic ventilatory response [279,280]. Likewise, the mechanisms underlying this plasticity have not yet been investigated. Although changes in metabolism, gas exchange and blood buffering do not appear to be involved in the altered ventilatory responses in adult quail [279], early postnatal CO2 exposure in rats causes changes in respiratory mechanics that persist into adulthood [281]. Therefore, it is currently not possible to determine whether changes in the hypercapnic response reflect changes in CO2 chemosensitivity, CNS integration or respiratory mechanics. However, since the hypoxic response was not affected by neonatal hypercapnia in rats [277], the role of the observed changes in respiratory mechanics in the blunted hypercapnic response following developmental hypercapnia is questionable. F.
Metabolism
Changes in metabolic rate are typically matched by corresponding increases or decreases in ventilation [282]. Sant’Anna and Mortola [283,284] investigated whether chronic alterations in metabolic rate induce developmental plasticity in respiratory control. In the first set of experiments, rats were raised for the first three postnatal weeks in small (6 pups/litter) or large (16 pups/litter) litter sizes [283]. Large-litter rats had reduced growth and lower metabolic rates prior to weaning (presumably reflecting reduced caloric intake due to increased food competition). Ten days later, resting ventilation was greater in large-litter vs. small-litter rats. However, the increased ventilation was matched by a greater mass-specific metabolic rate (i.e., the ventilation to metabolism ratio was unchanged). Hypoxic and hypercapnic ventilatory responses were unaffected by developmental litter size. In a second study [284], rats were exposed to cold (14 C) for the first three postnatal weeks, resulting in a chronic elevation of metabolic rate during this period. Aside from a small change toward a slower and deeper breathing pattern, no changes in resting ventilation or metabolism or in the hypoxic and hypercapnic ventilatory responses were detected in cold-reared rats when studied approximately 10 days later. These studies suggest that chronic alterations in metabolic rate during development have minimal consequences for the respiratory control system. However, it is likely that the experimental treatments affected more than metabolic rate (e.g., stress hormones) and the data should be interpreted with caution. As reviewed in the preceding sections, developmental perturbations of respiratory gases may cause long-lasting changes in respiratory control. However, the effects of prolonged changes in ventilation induced by altered respiratory gases (i.e., increased respiratory drive) vs. effects of altered
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respiratory gases themselves (i.e., tissue PO2 and PCO2) are unknown. Since, chronic alterations in metabolic rate modify ventilation with minimal effects on blood gases (i.e., ventilation to metabolism ratio unchanged; [282], studies of developmental metabolic derangements may shed light on this issue [284]. Specifically, the studies by Sant’Anna and Mortola [283,284] suggest that the change in blood gases may be the stimulus for long-lasting plasticity following developmental hypoxia, hyperoxia, and hypercapnia. G. Sensory Denervation
Sensory denervation during development may induce plasticity in respiratory control. Carotid body denervation typically results in hypoventilation during normoxia and eliminates, or greatly reduces, ventilatory responses to hypoxia in neonatal and adult mammals [7]. In many mammalian species, both young and adult animals exhibit substantial plasticity following carotid denervation, spontaneously recovering resting blood gases and hypoxic ventilatory responsiveness over a period of weeks to years, although this recovery is somewhat less in humans (reviewed in Ref. [7]). Studies in rats, pigs and goats suggest that this recovery may occur sooner or more completely if the carotid body denervation occurs in early postnatal life [7,44,45,285]. In adults, recovery following carotid denervation may involve an upregulation of aortic body chemoreceptor function, abdominal and/or central hypoxia-sensitive tissues, or chemosensory pathways [286–291]. In contrast, functional, O2-sensitive aortic chemoreceptors may already be present during the neonatal period. Aortic chemosensitivity (assessed by ventilatory responses to local NaCN injection) is normally lost at around 8 days of age in piglets, but remains prominent in piglets denervated neonatally [45,285]; similar observations have been reported for rats [44]. Moreover, there appears to be a critical period during development (around the second week) when piglets are more likely to exhibit irregular breathing and apneas following carotid denervation [292]; denervation before or after this critical period has no effect on respiratory stability. The relatively normal breathing observed following denervation prior to this critical period may reflect the immediate availability of functional aortic chemoreceptors [285]. Together, these data suggest that neonatal animals are able to capitalize on redundancy in the respiratory control system that is normally lost during maturation, and this capacity may contribute to age-dependent differences in recovery from carotid denervation. Given the dramatic recovery of hypoxic ventilatory responses following carotid denervation, it is interesting that hypoxic responses are still reduced at 3 months of age in rats exposed to developmental hyperoxia, and may remain so permanently [251]. Indeed, residual hypoxic sensitivity is
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unaffected by vagotomy in these rats and is abolished following bilateral section of the CSN [249], suggesting that aortic chemoreceptors do not upregulate in this model. It is possible that this difference between carotid denervation vs. developmental hyperoxia reflects the impairment of both carotid and aortic chemoreceptors by hyperoxia in the latter case. Alternatively, aortic chemoreceptor upregulation may be inhibited in hyperoxia-treated rats. Aortic chemosensitivity following carotid body denervation is serotonin-dependent [45], and serotonin and serotonin receptors are upregulated in the chemosensitive portion of the aorta following bilateral carotid denervation in neonatal rats and piglets [293]. Since unilateral carotid denervation elicits no change in serotonin or serotonin receptors (unpublished data cited in Ref. [293]) the residual carotid body function in hyperoxia-treated rats may be sufficient to prevent compensatory aortic chemosensitivity. H. Neonatal Maternal Separation
Stressful conditions, such as restraint or immobilization, alter ventilatory control in adult mammals [294–296]. These effects may persist at least 24 h after immobilization has ended, indicating plasticity in ventilatory control mechanisms [296]. Recent evidence indicates that stress during early postnatal development caused by maternal separation has lasting effects on the control of breathing as well. In these studies, rat pups were isolated from their mothers for 3 h/day on 10 consecutive days, beginning on postnatal day 3 [297,298]. As adults, rats that experienced neonatal maternal separation exhibited reduced hypercapnic ventilatory responses, possibly related to changes in the paraventricular nucleus of the hypothalamus [297]. Although a decreased hypercapnic response is also observed following immobilization stress in adult rats [296], the enduring nature of this plasticity suggests that these effects could be unique to development. Moreover, neonatal maternal separation enhances adult hypoxic ventilatory and phrenic responses in male (but not female) rats ([298]; Kinkead, R., personal communication). In contrast, hypoxic responses were unaffected by immobilization stress in adult male rats [296]. It is unknown whether neonatal maternal separation and adult immobilization produce comparable stress responses, or to what extent the duration of exposures (10 days in neonates, 1–2 days in adults) alters this plasticity. However, these studies suggest the existence of critical periods for stress-related respiratory plasticity. It is important to note that altered respiratory gases can induce stress responses (e.g., stress hormone secretion), although this responsiveness may be relatively low during development [299–301]. Thus, activation of stress responses during development may be one mechanism by which altered respiratory gases induce developmental plasticity. However, there is no
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direct evidence to support a causal role for a generalized stress response in the examples of developmental plasticity (i.e., hypoxia, hyperoxia, and hypercapnia) described in the preceding sections. Indeed, the variability in plasticity (magnitude, direction, and altered reflex) and underlying mechanisms (e.g., chemosensitivity vs. respiratory mechanics) argue against a single mechanism for developmental plasticity in respiratory control. IV.
Sex Hormones
Sex is emerging as an important consideration in the study of developmental plasticity. Several examples of developmental plasticity are specific to either males or females. Examples include plasticity induced by neonatal hypoxia [233,302] and maternal separation [298] in rats, and embryonic hypercapnia in birds [279]. Other forms of developmental plasticity are sex insensitive (e.g., developmental hyperoxia; [251]), but most models have not been studied with respect to sex. This is an important consideration as sex may impact the assessment of plasticity, and the value of therapeutic strategies intended to induce plasticity may be sex-specific. Future studies will benefit by explicitly considering sex in their experimental designs. The mechanisms by which sex interacts with plasticity are unknown, although hormones are likely to be involved in both developmental and adult neuroplasticity [303–305]. Sex hormones are believed to influence the control of breathing [306] and have recently been implicated in adult respiratory plasticity following intermittent hypoxia (i.e., LTF, [109,110]). Moreover, manipulation of sex hormones during development can in itself influence adult respiratory control. Male and female adult rats exhibit distinct ventilatory responses to injection of aspartic acid, an NMDA receptor agonist [307,308]. Neonatal treatment with sex hormones (testosterone in females or estradiol in males) can switch the female ventilatory response into the male-type response, and vice versa, possibly by altering NMDA receptor expression [308–310]; these effects of neonatal hormone treatment do not appear until after sexual maturity [308]. In addition, male rats treated with estradiol at 5 days of age had reduced hypercapnic ventilatory responses as adults despite similar resting ventilation [310]. Consequently, environmental factors that alter perinatal hormone levels (e.g., maternal stress in rats; [311]), could have long-term effects on the control of breathing. V.
Spinal Cord Injury (SCI) and Respiratory Plasticity
One strategy for promoting functional motor recovery following incomplete SCI is to strengthen (and better utilize) existing, non-injured pathways [312].
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The inherent plasticity of the respiratory neural control system makes this approach a viable therapeutic option for improving respiratory muscle control following SCI. Respiratory deficits following cervical SCI are determined by the severity and segmental level of the injury. High cervical SCI (lesions rostral to C4) are immediately life threatening because inspiratory premotor drive to spinal respiratory motoneurons is interrupted. Thus, high cervical lesions generally require immediate and sustained ventilatory support, whereas spinal lesions caudal to C4 produce comparatively less respiratory dysfunction. However, even low cervical and thoracic SCI patients are at risk for respiratory-related disorders (e.g., pneumonia; [313]) and have impaired airway protective reflexes (e.g., cough; [314]). Early spontaneous recovery of motor function following SCI is often attributed to resolution of spinal shock, inflammation and edema in the spinal cord [315]. Subsequent improvements may reflect increased respiratory muscle strength [316], altered pulmonary mechanics [317], and recruitment of accessory inspiratory muscles not normally used to generate airflow [316,318]. An additional possibility is that time-dependent neuroplasticity in surviving neurons/networks improves respiratory motor function in the months to years following SCI. Much evidence suggests that neuroplasticity can improve respiratory motor function in animal models of SCI (reviewed below). A recent case report provides hope that the same is true for human cervical SCI patients [319]. A C2 injured, ventilator-dependent patient with complete loss of motor and sensory function (grade A; American Spinal Injury Association) experienced no significant motor recovery in the first five years post-injury, although an MRI indicated that up to 20% of C2 spinal tissue was intact. The patient then underwent a rigorous activity-based recovery program. Over years five to eight post-injury, the patient demonstrated substantial motor recovery (two ASIA grades) that was attributed to the training. EMG studies suggest that the patient may have regained some voluntary control of the diaphragm. The significance of this respiratory motor recovery is evident in the following quotation from the patient, A ventilator failure . . . would have been a terrifying experience because I couldn’t really breathe. Now, I can breathe quite well . . . I am able to move my diaphragm, an ability that was achieved by exercise and training [319]. This delayed motor recovery probably reflects plasticity in the small amount of spared respiratory pathways. Several laboratories are currently investigating if and how neuroplasticity can enhance respiratory function following SCI in animal models. In this section, we will review respiratory neural control and plasticity following injuries to the spinal cord including hemisection, contusion, and chronic deafferentation via dorsal rhizotomy.
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A. Spinal Hemisection
Animal models of respiratory dysfunction following SCI represent a compromise between what is seen clinically, what is survivable by laboratory animals without extensive respiratory support, and a clearly definable lesion. The most common experimental lesion used to study respiratory neural control following SCI is cervical hemisection (from midline to the lateral edge of the cord) rostral to the phrenic motor nucleus (for review see Refs. [8,320]). Cervical hemisection is reproducible, quantifiable and has a low mortality rate in rats. The major advantage of the hemisection model is that it permits study of existing, but normally ineffective, spinal pathways that cross the spinal midline caudal to the injury and project to phrenic motoneurons (reviewed in detail below). However, a limitation of spinal hemisection as an SCI model is that it is not representative of the most common clinical injuries (i.e., spinal contusion; [312]). Minute ventilation is sufficient to maintain arterial blood gases during normoxia within a few days following C2 hemisection [321], but hypoventilation may occur in the first 1–2 h post-injury (Fuller, D.D., Golder, F.J. and Mitchell, G.S., unpublished observations). Respiratory insufficiency following C2 hemisection may also be revealed during anesthesia [322,323] or hypoxic or hypercapnic challenge. Rats with chronic C2 hemisection breathe with increased frequency and decreased tidal volume at 1 and 2 months post-injury when anesthetized [323] or awake (Fuller, D.D. and Mitchell, G.S., unpublished observations). Tidal volume continues to diminish over a period of weeks post-C2 hemisection, but breathing frequency progressively increases resulting in a constant minute ventilation (Fuller, D.D. and Mitchell, G.S., unpublished observations). The mechanisms producing the alterations in breathing pattern following C2 hemisection are not fully known, but partly result from pulmonary vagal feedback [323]. In addition, the altered breathing pattern may reflect supraspinal plasticity [324] or changes in pulmonary mechanics (e.g., reduced chest wall compliance; [317]). During breathing at rest, the phrenic nerve or hemidiaphragm ipsilateral to C2 hemisection is silent (i.e., no inspiratory bursting) in the days to weeks following the injury [30,321,323–325]. However, chemical or pharmacological stimulation of respiratory drive produces rhythmic inspiratory phrenic nerve activity below the hemisection [8]. Inspiratory bursts occur at the same frequency as those in the contralateral phrenic nerve, but amplitude is considerably less [30,322–324]. How does respiratory activity recur despite elimination of all ipsilateral bulbospinal inputs? In rats, bulbospinal respiratory neurons project axons bilaterally to phrenic motoneurons [326–328]. Although most crossed pathways decussate in the brainstem, an apparently ineffective synaptic pathway
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crosses the spinal midline in rats caudal to C2 [326]. Phrenic motor output caudal and ipsilateral to cervical hemisection results from activation of these existing but previously ineffective synaptic pathways to phrenic motoneurons [8]. This effect has been termed the crossed phrenic phenomenon [8,329]. Many published studies of the crossed phrenic phenomenon do not use histological techniques to document that all descending ipsilateral tracts have been severed. Accordingly, the existence of a crossed-spinal pathway is not required to explain the crossed phrenic phenomenon in all published studies. One alternative neural pathway to phrenic motoneurons could be dendrites from contralateral motoneurons that cross the spinal midline as originally suggested by Ref. [330]. Phrenic motoneuron dendrites that cross the spinal midline are more prevalent in neonatal rats [331,332], but still may be present in adult animals [332]. A crosseddendritic pathway could transmit inspiratory impulses to motoneurons below the hemisection. Regardless of the anatomical substrate, the crossed phrenic phenomenon is one of the best examples of motor recovery following SCI and it provides an opportunity to examine the mechanisms underlying recruitment and plasticity of functionally latent synapses [333]. The crossed phrenic phenomenon occurs in a wide range of species including rats [30,321,323,324], rabbits [334,329], dogs [329,335], cats [335,336], guinea pigs [333] and woodchucks [329]. However, Rosenbluth and Ortiz [329] did not observe the crossed phrenic phenomenon in 3 of 4 acutely hemisected monkeys. In the fourth monkey, cervical spinal hemisection did not paralyze the ipsilateral diaphragm, but ispilateral hemidiaphragm contractions were abolished by cutting the contralateral phrenic nerve [329]. Rosenbluth and Oritz concluded that diaphragm contractions were produced by impulses traveling in axons of the phrenic nerve contralateral to hemisection, and therefore did not represent crossed phrenic activity. However, the post-hemisection time interval before crossed phrenic pathways can be recruited is variable between species (an observation that may account for the failure of Rosenbluth and Ortiz [329] to find crossed phrenic activity in primates). The crossed phrenic phenomenon can be induced immediately post-hemisection in cats, dogs, rabbits and woodchucks [329]. In contrast, young rats and guinea pigs require a delay ranging from 3 to 24 h before chemical stimulation will reveal crossed phrenic pathways [337,338]. However, crossed phrenic pathways can be activated immediately following hemisection if rats are treated with the serotonin precursor 5-hydroxytryptophan (5-HTP) [30,339]. Thus, following acute hemisection in rats, crossed phrenic pathways are present, but functionally latent [8]. One possibility is that activation of crossed phrenic pathways is prevented by inhibitory influences associated with acute SCI (e.g., hemorrhage, swelling, etc.; [312]). A more likely
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scenario is that activation of crossed phrenic pathways requires necessary preconditions (e.g., removal of active inhibition, [321]; morphological plasticity, [8]; insertion of glutamate receptors into post-synaptic membranes, [135]) not present immediately post-hemisection [30]. Consistent with this interpretation, robust crossed phrenic responses occur in acutely C2 hemisected rats given a pre-conditioning lesion CDR but not in control or sham-operated rats [340]. CDR may create necessary preconditions by enhancing serotonergic terminal density [128] or neurotrophin concentration in the cervical spinal cord [341]. Both of these putative changes could increase efficacy in existing spinal synapses (see section above on Brief intermittent hypoxia). The crossed phrenic phenomenon is influenced by age of the animal at the time of injury [338,342,342a]. Older rats require no delay between hemisection injury and recruitment of crossed phrenic pathways during asphyxia [338]. Goshgarian [338] speculates that crossed phrenic pathways mature during the normal aging process, resulting in more effective synaptic transmission. However, this hypothesis may not be true of all synaptic inputs to phrenic motoneurons. For example, long-latency (41.0 ms) crossed phrenic potentials evoked by spinal cord stimulation are revealed following systemic treatment with 5-HTP [339]. These long-latency potentials are present in young (3–5 month) but not old (1.5–2 years) rats [342]. Thus, age-related developmental plasticity of crossed phrenic pathways is a complex process that requires further investigation. Sex-related influences on crossed phrenic pathways have not been specifically investigated. Separate laboratories studying exclusively male or female rats following C2 hemisection have reported qualitatively similar data during chemical or pharmacological induction of crossed phrenic activity [8,30,323,324]. However, the rate of spontaneous recovery of inspiratory motor activity in rats may be more rapid in males [30]. Male rats also appear to recover locomotor function more rapidly than female rats following C2 hemisection (Golder, F.J., personal communication). Further, sex has a critical influence on spinal respiratory plasticity in the form of phrenic LTF [83,84,109,110]. Genetic differences in crossed phrenic expression are also unknown. Genetics are suspected to influence spinal respiratory plasticity [90] and motor recovery following SCI [343]. Moreover, deficits in inspiratory tidal volume following C2 hemisection are different between rat strains (Fuller, D.D. and Mitchell, G.S., unpublished observations). Collectively, these observations suggest that sex and genetic influences on crossed phrenic activity merit further investigation. Although the crossed phrenic phenomenon has been an important model of respiratory motor recovery following SCI for over 100 years, it was not known whether crossed phrenic activity made a functionally
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meaningful contribution to inspiratory tidal volume. By comparing ventilation between rats with either C2 hemisection alone or C2 hemisection þ ipsilateral phrenicotomy (i.e., preventing crossed phrenic activity from reaching the diaphragm), Golder and colleagues [322] assessed the contribution of these crossed pathways to breathing in rats. Ipsilateral phrenicotomy did not alter the rapid shallow breathing pattern in C2 injured rats. However, the ability to generate large inspiratory volumes (e.g., during augmented breaths) was significantly impaired if crossed phrenic activity was prevented. Thus, crossed phrenic activity is physiologically significant in terms of (large) tidal volume generation [322]. B. Serotonin and Crossed Phrenic Pathways
Serotonin may be a key element in the spontaneous improvement of locomotor and respiratory function following SCI [131,324,344–349]. Following pre-treatment with the serotonin synthesis inhibitor parachlorophenylalanine (p-CPA), the crossed phrenic phenomenon is observed in fewer rats, and the magnitude of phrenic activation on the hemisected side is reduced [350]. Similarly, treatment with the serotonergic neurotoxin 5,7 dihydroxytryptamine prior to injury significantly reduces crossed phrenic activity 2 months following hemisection [324]. Conversely, the serotonin precursor 5-HTP activates crossed phrenic pathways in spinally hemisected rats [30,339]. Recruitment of crossed phrenic pathways following 5-HTP is reversed following serotonin receptor antagonism with methysergide [339,342,351,352]. The specific serotonin receptor subtype necessary to recruit crossed phrenic pathways appears to be the 5-HT2A receptor [348,131]. 5-HT2A receptors located on phrenic motoneurons [131] may contribute to recruitment of crossed phrenic pathways. Further evidence for a spinal mechanism underlying crossed phrenic pathway activation comes from studies utilizing spinal cord electrical stimulation [30,339,342]. Shortly after i.v. 5-HTP, a short-latency (0.7 ms) crossed-phrenic-compound action potential is evoked by stimulating the ventrolateral spinal cord at C2 [339]. The latency of this response indicates that spinal cord neurons must be responsible, since this is insufficient time for a relay through supraspinal structures. C. Spontaneous Motor Recovery in Crossed Phrenic Pathways
Time-dependent plasticity occurs following cervical hemisection, strengthening crossed phrenic pathways [8]. For example, hours to days posthemisection in rats, inspiratory motor output ipsilateral to the injury can be induced only through increased ventilatory drive or by pharmacological
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means [8]. However, within two weeks, small crossed phrenic inspiratory bursts are observed at baseline conditions (Pa CO 2 40 mm Hg, PaO2 4 100 mm Hg) in approximately 50% of anesthetized rats ([30], Golder, F.J., personal communication; but see Ref. [325]). Spontaneous recovery grows stronger with time; one to two months post-C2 hemisection, spontaneous inspiratory bursts during quiet breathing (eupnea) are observed in the phrenic nerve or hemidiaphragm ipsilateral to injury in most C2 hemisected rats [324,325], although their amplitude remains small. The mechanisms underlying the spontaneous appearance of crossed phrenic activity are unknown, but may involve serotonin (see above). Initially, cervical hemisection decreases serotonin terminal density in the ipsilateral phrenic nucleus; however, over a period of days to weeks, serotonin terminal density increases to levels above that in normal rats [353]. Thus, descending serotonergic inputs project bilaterally to the phrenic nucleus in rats, which may provide a neuroanatomical basis for the timedependent recruitment of crossed phrenic pathways following chronic spinal hemisection. Spontaneous motor recovery in crossed phrenic pathways may develop in part secondary to morphological plasticity in the spinal cord following C2 hemisection [8,9]. For example, an increase in the number of dendro-dendritic appositions and synaptically active zones is observed in the ipsilateral phrenic motor nucleus a few hours post-C2 hemisection [354]. These rapid morphological effects are blunted when animals are given a serotonin synthesis inhibitor (para-chlorophenylalanine) prior to hemisection [355], consistent with effects of para-chlorophenylalanine on the crossed phrenic phenomenon [350]. The surface area of ipsilateral phrenic motoneuron somata are significantly decreased two weeks post-C2 hemisection, suggesting that motoneuron excitability increases following injury, a change that would enhance crossed phrenic activity [356]. D. Strengthening Crossed Phrenic Motor Output
One strategy for promoting functional recovery of respiratory motor output following cervical SCI is to strengthen (and better utilize) existing, non-injured pathways. This is a viable clinical strategy because the majority of SCIs are incomplete, and residual neural pathways are intact at the injury level [357]. Crossed phrenic pathway recruitment during heightened respiratory drive reflects ongoing recruitment or modulation (vs. plasticity) of an existing motor pathway since this effect is not persistent. However, chronic treatments may induce plasticity, strengthening crossed spinal synaptic inputs to phrenic motoneurons [30,340,358,359].
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Chronic pharmacological stimulation of respiratory drive with the adenosine receptor agonist theophylline appears to strengthen crossed phrenic pathways [358]. Following chronic (28 days) SCI theophylline treatment, 29 of 32 rats showed enhancement of crossed phrenic motor output. However, animals were studied shortly after theophylline treatment was terminated, raising the possibility that the motor recovery could reflect continued theophylline effects. The effects of theophylline were mediated by both adenosine A1 and serotonin 5HT2 receptors, possibly located on spinal motoneurons [348]. Our laboratory recently found that exposing C2 hemisected rats to CIH increased the efficacy of crossed phrenic pathways (Figure 6.4; [30]). C2 hemisected rats were exposed to CIH (5 min 11% O2, 5-min normoxia; 12 h/night) on day 7–14 post-injury, and were then studied in acute neurophysiological experiments. CIH-treated rats had substantially greater A
Spontaneous
B
Baseline Stimulating electrode
Evoked
Hypoxia
Control Hemisection
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Chronic Intermittent Hypoxia
Phrenic Recording
Figure 6.4 Chronic intermittent hypoxia strengthens normally ineffective pathways to phrenic motoneurons. The left panel (A) depicts crossed phrenic pathways to phrenic motoneurons (Data from Refs. 326, 376, and 377). Bulbospinal respiratory neurons have cell bodies in the medulla (ventral respiratory group) and project bilaterally to phrenic motoneurons. Bulbospinal projections which cross the spinal midline in the cervical spinal cord are known as crossed phrenic pathways. The terms ipsilateral and contralateral are generally used relative to the hemisection. Panel B depicts spontaneous inspiratory activity and compound action potentials (evoked via ventrolateral funiculus electrical stimulation) recorded in the phrenic nerve ipsilateral to C2 hemisection. Following chronic (2 week) C2 hemisection, ipsilateral phrenic inspiratory burst amplitude was significantly greater at baseline and during hypoxia in CIH-treated vs. normoxia-treated rats. Phrenic burst frequency was not different between groups at any time point. The panel on the far right shows crossed phrenic potentials evoked by 1000 mA stimulation current. Whereas the normoxia-treated rat has only a small evoked potential, the rat conditioned with CIH following chronic hemisection displays a clear evoked potential in the ipsilateral phrenic nerve suggesting that crossed phrenic pathways are enhanced by a spinal mechanism following CIH (arrow ¼ stimulus artifact) (figure modified from Ref. 30).
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inspiratory phrenic activity below hemisection at baseline and during chemoreceptor activation. In addition, short-latency (0.5–0.7 ms), spinally-evoked potentials recorded in the phrenic nerve ipsilateral to hemisection were of greater amplitude following CIH (Figure 6.4). These data strongly suggest increased efficacy of a monosynaptic spinal synapse [30]. Interestingly, pre-treatment (i.e., before C2 hemisection) with CIH had no discernable effect on crossed phrenic motor activity. Therefore, CIH-induced plasticity of crossed phrenic pathways requires necessary preconditions created by SCI. How might existing crossed phrenic activity be enhanced by CIH or other treatments? Our working hypothesis is that augmented crossed phrenic inspiratory burst amplitude following CIH represents increased synaptic strength of pre-existing crossed spinal pathways to phrenic motoneurons. Transformation of ineffective or silent synapses to functionally effective synaptic connections has considerable precedent [352]. For example, silent glutamatergic synapses in the rat dorsal horn are transformed into functional synapses by serotonin [360]. Serotonin may be of particular importance as CIH enhances phrenic motor output via serotonin-dependent mechanisms in spinal-intact rats [93] and can modulate crossed phrenic pathways [339,342]. Neurotrophins such as BDNF have the potential to strengthen existing synapses to phrenic motoneurons and may be involved in enhancing crossed phrenic activity. BDNF is critical for some forms of neuroplasticity (e.g., hippocampal long-term potentiation, [136]), can enhance neurotransmission at glutamatergic synapses [361] and, importantly, is upregulated in the cervical spinal cord following brief episodic hypoxia [362]. Correlative evidence provides further support for a role of BDNF in enhancing crossed phrenic pathways: a pre-conditioning lesion (CDR) increases BDNF protein levels in the ventral spinal cord [341] and also enhances crossed phrenic pathways [340]. However, preliminary data from our laboratory indicate that neither BDNF nor serotonin are significantly altered in the ventral cervical spinal gray matter two weeks following C2 hemisection, either with or without CIH-treatment [363]. This observation does not rule out a role for these molecules in CIH-induced spinal plasticity, but limits potential involvement to initiation vs. maintenance of increased synaptic strength. For example, in a number of other models of serotonin-dependent neuroplasticity, serotonin receptor activation is necessary to initiate, but not maintain the plasticity [117,364,365]; neurotrophins such as BDNF may play a similar role. Consistent with this idea, BDNF and receptor tyrosine kinase B mRNA expression transiently increase following C2 hemisection, but return to control levels within two weeks [356].
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Contralateral and Supraspinal Effects of Cervical Hemisection
Unilateral SCI often induces plasticity in contralateral motor pools. For example, after thoracic spinal hemisection, perinatal chicks regain posture and locomotor skills via increased reliance on contralateral motor activity [366]. Similarly, rats with cervical or thoracic hemisection injuries have altered locomotor patterns with increased reliance on contralateral motoneurons [367]. In contrast, contralateral respiratory motor output may be inhibited under certain conditions following unilateral spinal cord injury [324]. Two months post-hemisection, anesthetized, paralyzed, and ventilated rats have a decreased inspiratory burst frequency during quiet breathing and reduced contralateral phrenic burst amplitude during hypercapnia [324]. These adaptations may reflect supraspinal (e.g., medullary) plasticity because hypoglossal motor output is also attenuated following cervical spinal hemisection [324]. This diminution of contralateral respiratory motor output following a unilateral lesion is difficult to explain, particularly because chronic C2 hemisected rats maintain adequate ventilation ([321]; Fuller, D.D. and Mitchell, G.S., unpublished observations). However, rats that do not spontaneously express the crossed phrenic phenomenon do not exhibit reductions in contralateral phrenic output [320]. Further, rats receiving a dual injury (C2 hemisection and ipsilateral phrenicotomy) that prevents crossed phrenic motor output from reaching the diaphragm also do not show reductions in contralateral phrenic output [322]. Indeed, the contralateral phrenic nerve actually shows enhanced inspiratory burst amplitude during chemoreceptor stimulation in dual injury rats [322]. Thus, contralateral phrenic motor plasticity is critically influenced by the presence (or absence) of crossed phrenic activity. Compensatory plasticity in contralateral motor pools (i.e., greater motor output) may require necessary preconditions that are established only when crossed phrenic activity is prevented. F.
Cervical Contusion Injuries
Respiratory motor plasticity following cervical spinal contusion injuries is largely unexplored. The clinical literature documents recovery of respiratory motor function following cervical contusion (particularly with low cervical injury), but it is unknown if this recovery reflects neuroplasticity, a change in pulmonary mechanics, or other factors. El-Bohy et al. [368] demonstrated the feasibility of cervical contusion in rats as a model of respiratory dysfunction following SCI. Rats were able to survive a lateralized contusion at C2 created with the New York University contusion system (impact height ¼ 12.5 mm). Phrenic nerve recordings five weeks post-injury suggested a reduced dynamic range of phrenic motor output ipsilateral to the contusion. In specific, baseline motor output averaged almost 80% of
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that observed during asphyxia [368]. Normalization concerns make these data difficult to interpret and additional studies are needed. Following T8 contusion injury in rats (weight drop device, [369]), inspiratory tidal volume is reduced, breathing frequency is elevated, and minute ventilation is unchanged [370]. However, these changes in the pattern of breathing do not persist beyond the first week post-injury [370]. Interestingly, a single spinal injection of basic fibroblast growth factor at the time of injury prevented the initial changes in breathing pattern [370]. This report highlights the potential use of breathing as a functional outcome measure to assess experimental therapeutic approaches following SCI [371]. G. Dorsal Rhizotomy
Acute CDR reveals the crossed phrenic phenomenon in female rats, suggesting a contralateral afferent pathway that may inhibit crossed pathways to phrenic motoneurons [321]. Thus, crossed phrenic motor output following acute CDR may reflect removal of inhibitory afferent projections to phrenic motoneurons via polysynaptic pathways (reviewed in Ref. [372]). This effect may be species specific, as Rosenbluth and Ortiz [329] reported no effect of acute CDR on crossed phrenic activity in cats and rabbits. Chronic dorsal rhizotomy elicits morphological and neurochemical plasticity within the spinal cord. Chronic CDR increases serotonin terminal density in the immediate vicinity of labeled phrenic motoneurons and also increases phrenic motoneuron size [128]. Chronic dorsal rhizotomy also increases serotonin terminal density in the spinal dorsal horn [349,373,374]. Chronic CDR is associated with increased ventral cervical spinal concentrations of BDNF and a related neurotrophic factor, neurotrophin-3 [341]. In association, CDR has little impact on the shortterm hypoxic phrenic response and PHFD [128]. However, serotonindependent phrenic LTF is enhanced following chronic CDR [128]. The enhanced LTF is blocked by the 5-HT2 receptor antagonist ketanserin, indicating that this form of metaplasticity (i.e., enhanced LTF) exhibited normal serotonin receptor dependence [128]. Chronic CDR also reveals normally ineffective crossed phrenic pathways [340]. In acutely C2 hemisected rats, crossed phrenic potentials were evoked at a lower current in rats with chronic CDR [340]. Importantly, the phrenic nerves were sectioned prior to recording in these experiments in both control and CDR rats; thus inhibitory phrenic afferent inputs were absent in both experimental groups. In contrast to the LTF data (see above), serotonin receptor antagonism had minimal effect on crossed phrenic activity following chronic CDR [340]. Thus, serotonin receptor activation is not necessary to maintain enhanced crossed phrenic pathways following CDR,
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although this does not rule out a potential role for serotonin in initiating spinal plasticity. It is tempting to speculate that changes in spinal neurotrophins following CDR [341] contribute to enhancement of crossed phrenic pathways, but this hypothesis requires rigorous evaluation. Alternatively, respiratory plasticity following spinal sensory denervation does not necessarily result from a spinal mechanism. CDR can induce plasticity distant to the surgically affected area. For example, CDR (C3–C5) enhances LTF of hypoglossal motor output [375], even though the hypoglossal motor nucleus is rostral to the site of denervation. Therefore, changes in respiratory motor output following chronic dorsal rhizotomy may reflect plasticity at distant sites (see below). Bilateral thoracic (T2–T12) dorsal rhizotomy in goats induces functional and morphological plasticity [349]. After recovery from surgery, an intriguing pattern of severe ventilatory failure during even modest exercise, followed by progressive functional recovery, was observed [349]. In association with these functional deficits and/or recovery, increased serotonin concentrations and serotonin-immunoreactive terminal density were found in the thoracic and cervical spinal cord, suggesting that serotonin may be involved in the mechanism underlying functional recovery [349]. However, there was also evidence for increased spinal dopamine and norepinephrine concentrations [349]. One intriguing aspect of these changes in spinal neuromodulators is an observation that serotonin and dopamine increased at spinal segments C6–C7, the segments associated with the phrenic motor nucleus in goats [349]. One possible interpretation of these findings is that neurochemical changes outside the segments directly affected by surgery (and denervation) compensate for the loss of respiratory function and thereby contribute to functional recovery. These hypotheses remain to be tested. V.
Conclusion
The neurons and networks controlling ventilation express plasticity and metaplasticity in response to a myriad of stimuli including alterations in environmental gases and neural injury. Several unifying principles emerge from our review of respiratory plasticity. First, the pattern, duration, and severity of experimental or environmental stimulation is an important determinant of respiratory plasticity. Second, both sex and genetics must be considered when evaluating phenotypes associated with, or mechanisms responsible for, respiratory plasticity. Finally, development and aging exert considerable influence on the expression of respiratory neuroplasticity. Some forms of respiratory plasticity are unique to development; others may be unique to later periods (e.g., adult or geriatric animals).
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The physiological significance of respiratory plasticity is clear in some examples (e.g., recovery of ventilation following spinal injury) and perhaps less so in other cases. Through continued study we may come to appreciate the physiological significance of these and other forms of respiratory plasticity. For example, LTF of upper airway muscle activity may stabilize the pharyngeal airways during sleep, and a loss of LTF with age may contribute to OSA. Respiratory neurobiologists are beginning to unravel the mechanisms underlying certain forms of plasticity (e.g., LTF, VAH, etc.) and these efforts are leading to a greater understanding of respiratory neural control. Equally important, such studies may lead to insights concerning mechanisms of plasticity throughout the CNS. It is our hope that careful exploration of respiratory plasticity, using molecular through behavioral approaches in comparative and clinical studies, will lead to an understanding of the role played by plasticity (or lack of plasticity) in respiratory disorders. Further, an understanding of respiratory neuroplasticity mechanisms may provide the rationale to harness this plasticity for rehabilitative efforts (e.g., following SCI) or for the treatment of respiratory-related pathologies such as OSA. Acknowledgments D.D. Fuller was supported by a Parker B. Francis Fellowship in Pulmonary Research. R.W. Bavis was supported by NIH NRSA post-doctoral fellowship award HL70506. Work conducted in G.S. Mitchell’s laboratory was funded by the National Institutes of Health (HL/NS 69064, HL53319, and HL 65383). We thank Brad Hodgeman for preparation of the figures and Drs. T. Baker-Herman, F.J. Golder and S.M. Johnson for their helpful critique of the manuscript. References 1.
Mitchell, G.S. and Johnson, S.M., Neuroplasticity in respiratory motor control, J. Appl. Physiol. 94, 358–374, 2003. 2. Bliss, T.V.P. and Lomo, T., Long-lasting potentiation of synaptic transmission in the dentate area of anaesthetized rabbit following stimulation of the perforant path, J. Physiol. 232, 331–356, 1973. 3. McGaugh, J.L., Memory—A century of consolidation, Science 287, 248–251, 2000. 4. Mitchell, G.S., Baker, T.L., Nanda, S.A., Fuller, D.D., Zabka, A.G., Hodgeman, B.A., Bavis, R.W., Mack, K.J. and Olson, E.B., Jr., Intermittent hypoxia and respiratory plasticity, J. Appl. Physiol. 90, 2466–2475, 2001. 5. Feldman, J.L., Mitchell, G.S. and Nattie, E.E., Breathing: Rhythmicity, plasticity, chemosensitivity, Annu. Rev. Neurosci. 26, 239–266, 2003.
200 6. 7. 8.
9. 10.
11. 12. 13.
14. 15. 16.
17.
18. 19.
20. 21.
22.
23.
Fuller et al. Carroll, J.L., Developmental plasticity in respiratory control, J. Appl. Physiol. 94, 375–389, 2003. Forster, H.V., Plasticity in the control of breathing following sensory denervation, J. Appl. Physiol. 94, 784–794, 2003. Goshgarian, H.G., The crossed phrenic phenomenon: A model for plasticity in the respiratory pathways following spinal cord injury, J. Appl. Physiol. 94, 795–810, 2003. Mantilla, C. and Sieck, G., Mechanisms underlying motor unit plasticity in the respiratory system, J. Appl. Physiol. 94, 1230–1241, 2003. Morris, K.F., Baekey, D.M., Nuding, S.D., Dick, T.E., Shannon, R. and Lindsey, B.G., Neural network plasticity in respiratory control, J. Appl. Physiol. 94, 1242–1252, 2003. Abraham, W.C. and Bear, M.F., Metaplasticity: The plasticity of synaptic plasticity, Trends Neurosci. 19, 126–130, 1996. Abraham, W.C. and Tate, W.P., Metaplasticity: A new vista across the field of synaptic plasticity, Prog. Neurobiol. 52, 303–323, 1997. Katz, P.S. and Edwards, D.H., Metamodulation: The control and modulation of neuromodulation, in Beyond Neurotransmission: Neuromodulation and its Importance for Information Processing, Katz, P.S., ed., New York, Oxford University Press, pp. 349–381, 1999. Byrne, J.H., Synapses. Plastic plasticity, Nature 389, 791–792, 1997. Nattie, E.E., CO2, brainstem chemoreceptors and breathing, Prog. Neurobiol. 59, 299–331, 1999. Nattie, E.E. and Prabhakar, N.R., Peripheral and central chemosensitivity: Multiple mechanisms, multiple sites? A workshop summary, Adv. Exp. Med. Biol. 499, 73–80, 2001. Ling, L., Olson, E.B., Jr., Vidruk, E.H. and Mitchell, G.S., Developmental plasticity of the hypoxic ventilatory response, Respir. Physiol. 110, 261–268, 1997. Bisgard, G.E., Carotid body mechanisms in acclimatization to hypoxia, Respir. Physiol. 121, 237–246, 2000. Prabhakar, N.R. and Kline, D.D., Ventilatory changes during intermittent hypoxia: Importance of pattern and duration, High Alt. Med. Biol. 3, 195–204, 2002. Mifflin, S.W., Short-term potentiation of carotid sinus nerve inputs to neurons in the nucleus of the solitary tract, Respir. Physiol. 110, 229–236, 1997. Gozal, D., Gozal, E. and Simakajornboon, N., Signaling pathways of the acute hypoxic ventilatory response in the nucleus tractus solitarius, Respir. Physiol. 121, 209–221, 2000. Cheng, Z., Guo, S.Z., Lipton, A.J. and Gozal, D., Domoic acid lesions in nucleus of the solitary tract: Time-dependent recovery of hypoxic ventilatory response and peripheral afferent axonal plasticity, J. Neurosci. 22, 3215– 3226, 2002. Kinkead, R., Bach, K.B., Johnson, S.M., Hodgeman, B.A. and Mitchell, G.S., Plasticity in respiratory motor control: Intermittent hypoxia and hypercapnia activate opposing serotonergic and noradrenergic modulatory systems, Comp. Biochem. Physiol. A 130, 207–218, 2001.
Respiratory Neuroplasticity 24. 25. 26. 27.
28.
29.
30.
31. 32. 33. 34. 35. 36.
37.
38.
39.
40. 41.
201
Dick, T.E. and Coles, S.K., Ventrolateral pons mediates short-term depression of respiratory frequency after brief hypoxia, Respir. Physiol. 121, 87–100, 2000. Siniaia, M.S., Young, D.L. and Poon, C.S., Habituation and desensitization of the Hering-Breuer reflex in rat, J. Physiol. 523, 479–491, 2000. Blitz, D.M. and Ramirez, J.M., Long-term modulation of respiratory network activity following anoxia in vitro, J. Neurophysiol. 87, 2964–2971, 2002. Johnson, S.M., Wilkerson, J.E., Henderson, D.R., Wenninger, M.R. and Mitchell, G.S., Serotonin elicits long-lasting enhancement of rhythmic respiratory activity in turtle brain stems in vitro, J. Appl. Physiol., 91, 2703–2712, 2001. McCrimmon, D.R., Zuperku, E.J., Hayashi, F., Dogas, Z., Hinrichsen, C.F., Stuth, E.A., Tonkovic-Capin, M., Krolo, M. and Hopp, F.A., Modulation of the synaptic drive to respiratory premotor and motor neurons, Respir. Physiol. 110, 161–176, 1997. Hayashi, F., Hinrichsen, C.F. and McCrimmon, D.R., Short-term plasticity of descending synaptic input to phrenic motoneurons in rats, J. Appl. Physiol. 94, 1421–1430, 2003. Fuller, D.D., Johnson, S.M., Olson, E.B., Jr. and Mitchell, G.S., Synaptic pathways to phrenic motoneurons are enhanced by chronic intermittent hypoxia following cervical spinal cord injury, J. Neurosci. 23, 2993–3000, 2003. Bazzy, A.R., Effect of hypoxia on neuromuscular transmission in the developing diaphragm, J. Appl. Physiol. 76, 708–713, 1994. Orem, J. and Netick, A., Behavioral control of breathing in the cat, Brain Res. 366, 238–253, 1986. Orem, J., Behavioral inspiratory inhibition: Inactivated and activated respiratory cells, J. Neurophysiol. 62, 1069–1078, 1989. Gallego, J. and Gaultier, C., Respiratory behavior, Rev. Mal. Respir. 17, 41–49, 2000. Gallego, J., Nsegbe, E. and Durand, E., Learning in respiratory control, Behav. Modif. 25, 495–512, 2001. Durand, E., Dauger, S., Vardon, G., Gressens, P., Gaultier, C., De Schonen, S. and Gallego, J., Classical conditioning of breathing pattern after two acquisition trials in 2-day-old mice, J. Appl. Physiol. 94, 812–818, 2003. Bisgard, G.E. and Neubauer, J.A., Peripheral and central effects of hypoxia, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 617–668, 1995. Baker, T.L., Fuller, D.D., Zabka, A.G. and Mitchell, G.S., Respiratory plasticity: Differential actions of continuous and episodic hypoxia and hypercapnia, Respir. Physiol. 129, 25–35, 2001. Fitzgerald, R.S. and Lahiri, S., Reflex responses to chemoreceptor stimulation, in Handbook of Physiology: Control of Breathing, Cherniack, N.S. and Widdicombe, J.D., eds., Bethesda, American Physiological Society, pp. 313– 362, 1986, Brophy, S., Ford, T.W., Carey, M. and Jones, J.F., Activity of aortic chemoreceptors in the anaesthetized rat, J. Physiol. 514, 821–828, 1999. Neubauer, J.A., Melton, J.E. and Edelman, N.H., Modulation of respiration during brain hypoxia, J. Appl. Physiol. 68, 441–451, 1990.
202 42.
43.
44. 45.
46.
47. 48.
49. 50.
51.
52. 53.
54. 55.
56.
57.
58.
Fuller et al. Nolan, P.C., Dillon, G.H. and Waldrop, T.G., Central hypoxic chemoreceptors in the ventrolateral medulla and caudal hypothalamus, Adv. Exp. Med. Biol. 393, 261–266, 1995. Solomon, I.C., Excitation of phrenic and sympathetic output during acute hypoxia: Contribution of medullary oxygen detectors, Respir. Physiol. 121, 101–117, 2000. Serra, A., Brozoski, D., Hedin, N., Franciosi, R. and Forster, H.V., Mortality after carotid body denervation in rats, J. Appl. Physiol. 91, 1298–1306, 2001. Serra, A., Brozoski, D., Hodges, M., Roethle, S., Franciosi, R. and Forster, H.V., Effects of carotid and aortic chemoreceptor denervation in newborn piglets, J. Appl. Physiol. 92, 893–900, 2002. Morris, K.F., Arata, A., Shannon, R. and Lindsey, B.G., Long-term facilitation of phrenic nerve activity in cats: Responses and short time scale correlations of medullary neurones, J. Physiol. 490, 463–480, 1996. Lutherer, L.O. and Williams, J.L., Stimulating fastigial nucleus pressor region elicits patterned respiratory responses, Am. J. Physiol. 250, R418–R426, 1986. Xu, F. and Frazier, D.T., Respiratory-related neurons of the fastigial nucleus in response to chemical and mechanical challenges, J. Appl. Physiol. 82, 1177–1184, 1997. Powell, F.L. Milsom, W.K. and Mitchell, G.S., Time domains of the hypoxic ventilatory response, Respir. Physiol. 112, 123–134, 1998. Bianchi, A.L., Denavit-Saubie´, M. and Champagnat, J., Central control of breathing in mammals: Neuronal circuitry, membrane properties, and neurotransmitters, Physiol. Rev. 75, 1–45, 1995. Eldridge, F.L. and Millhorn, D.E., Oscillation, gating, and memory in the respiratory system, in Handbook of Physiology: Control of Breathing, Cherniack, N.S. and Widdicombe, J.D., eds, Bethesda, American Physiological Society, pp. 93–114, 1986. Wagner, P.G. and Eldridge, F.L., Development of short-term potentiation of respiration, Respir. Physiol. 83, 129–139, 1991. Georgopoulos, D., Bshouty, Z., Younes, M. and Anthonisen, N.R., Hypoxic exposure and activation of the afterdischarge mechanism in conscious humans, J. Appl. Physiol. 69, 1159–1164, 1990. Fregosi, R.F., Short-term potentiation of breathing in humans, J. Appl. Physiol. 71, 892–899, 1991. Engwall, M.J., Daristotle, L., Niu, W.Z., Dempsey, J.A. and Bisgard, G.E., Ventilatory afterdischarge in the awake goat, J. Appl. Physiol. 71, 1511–1517, 1991. Mitchell, G.S., Powell, F.L., Hopkins, S.R. and Milsom, W.K., Time domains of the hypoxic ventilatory response in awake ducks: Episodic and continuous hypoxia, Respir. Physiol. 124, 117–128, 2001. Han, F., Subramanian, S., Dick, T.E., Dreshaj, I.A. and Strohl, K.P., Ventilatory behavior after hypoxia in C57BL/6J and A/J mice, J. Appl. Physiol. 91, 1962–1970, 2001. Kline, D.D. and Prabhakar, N.R., Role of nitric oxide in short-term potentiation and long-term facilitation: Involvement of NO in breathing stability, Adv. Exp. Med. Biol. 499, 215–219, 2001.
Respiratory Neuroplasticity 59.
60.
61.
62.
63.
64.
65.
66.
66a.
67. 68. 69.
70.
71.
72.
203
Kline, D.D., Overholt, J.L. and Prabhakar, N.R., Mutant mice deficient in NOS-1 exhibit attenuated long-term facilitation and short-term potentiation in breathing, J. Physiol. 539, 309–315, 2002. Xi, L., Smith, C.A., Saupe, K.W. and Dempsey, J.A., Effects of memory from vagal feedback on short-term potentiation of ventilation in conscious dogs, J. Physiol. 462, 547–561, 1993. Hayashi, F., Coles, S.K., Bach, K.B., Mitchell, G.S. and McCrimmon, D.R., Time-dependent phrenic nerve responses to carotid afferent activation: Intact vs. decerebellate rats, Am. J. Physiol. 265, R811– R819, 1993. Poon, C.S., Siniaia, M.S., Young, D.L. and Eldridge, F.L., Short-term potentiation of carotid chemoreflex: An NMDAR-dependent neural integrator, Neuroreport 10, 2261–2265, 1999. Young, D.L., Eldridge, F.L. and Poon, C.S., Integration–differentiation and gating of carotid afferent traffic that shapes the respiratory pattern, J. Appl. Physiol. 94, 1213–1229, 2003. Georgopoulus, D., Giannouli, E., Tsara, V., Argiropoulou, P., Patakas, D. and Anthonisen, N.R., Respiratory short-term poststimulus potentiation (after-discharge) in patients with obstructive sleep apnea, Am. Rev. Respir. Dis. 146, 1250–1255, 1992. Ahmed, M., Serrette, C., Kryger, M.H. and Anthonisen, N.R., Ventilatory instability in patients with congestive heart failure and nocturnal CheyneStokes breathing, Sleep 17, 527–534, 1994. Dahan, A., Berkenbosch, A., DeGoede, J., van den Elsen, M., Olievier, I. and van Kleef, J., Influence of hypoxic duration and posthypoxic inspired O2 concentration on short term potentiation of breathing in humans, J. Physiol. 488, 803–813, 1995. Menendez, A.A., Nuckton, T.J., Torres, J.E. and Gozal, D., Short-term potentiation of ventilation after different levels of hypoxia, J. Appl. Physiol. 86, 1478–1482, 1999. Eldridge, F.L., Post-hyperventilatory breathing: Different effects of active and passive hyperventilation, J. Appl. Physiol. 34, 422–430, 1973. Millhorn, D.E., Eldridge, F.L. and Waldrop, T.G., Pharmacologic study of respiratory afterdischarge, J. Appl. Physiol. 50, 239–244, 1981. Kline, D.D., Yang, T., Huang, P.L. and Prabhakar, N.R., Altered respiratory responses to hypoxia in mutant mice deficient in neuronal nitric oxide synthase, J. Physiol. 511, 273–287, 1998. Lipton, A.J., Johnson, M.A., Macdonald, T., Lieberman, M.W., Gozal, D. and Gaston, B., S-nitrosothiols signal the ventilatory response to hypoxia, Nature 413, 171–174, 2001. Gozal, D., Xue, Y.D. and Simakajornboon, N., Hypoxia induces c-Fos protein expression in NMDA but not AMPA glutamate receptor labeled neurons within the nucleus tractus solitarii of the conscious rat, Neurosci. Lett. 262, 93–96, 1999. Johnson, S.M. and Mitchell, G.S., Activity-dependent plasticity of descending synaptic inputs to spinal motoneurons in an in vitro turtle brainstem-spinal cord preparation, J. Neurosci. 20, 3487–3495, 2000.
204 73.
74.
75.
76. 77.
78.
79.
80.
80a.
81. 82.
83.
84.
85.
86.
87.
Fuller et al. Johnson, S.M. and Mitchell, G.S., Activity-dependent plasticity in descending synaptic inputs to respiratory spinal motoneurons, Respir. Physiol. Neurobiol. 131, 79–90, 2002. Jiang, C., Mitchell, G.S. and Lipski, J., Prolonged augmentation of respiratory discharge in hypoglossal motoneurons following superior laryngeal nerve stimulation, Brain Res. 538, 215–225, 1991. Bisgard, G.E. and Forster, H.V., Ventilatory responses to acute and chronic hypoxia, in Handbook of Physiology: Environmental Physiology, Fregly, M.J. and Blatteis, C.M., eds., pp. 1207–1239, 1996. Easton, P.A. and Anthonisen, N.R., Ventilatory response to sustained hypoxia after pretreatment with aminophylline, J. Appl. Physiol. 64, 1445–1450, 1988. Gershan, W.M., Forster, H.V., Lowry, T.F. and Garber, A.K., Effect of theophylline on ventilatory roll-off during hypoxia in goats, Respir. Physiol. 103, 157–164, 1996. Tatsumi, K., Pickett, C.K. and Weil, J.V., Effects of haloperidol and domperidone on ventilatory roll off during sustained hypoxia in cats, J. Appl. Physiol. 72, 1945–1952, 1992. Hoop, B., Beagle, J.L., Maher, T.J. and Kazemi, H., Brainstem amino acid neurotransmitters and hypoxic ventilatory response, Respir. Physiol. 118, 117–129, 1999. Gozal, D. and Gozal, E., Hypoxic ventilatory roll-off is associated with decreases in protein kinase C activation within the nucleus tractus solitarius of the rat, Brain Res. 774, 246–249, 1997. Gozal, D., Simakajornboon, N., Czapla, M.A., Xue, Y.D., Gozal, E., Vlasic, V., Lasky, J.A. and Liu, J.Y., Brainstem activation of platelet-derived growth factor-beta receptor modulates the late phase of the hypoxic ventilatory response, J. Neurochem. 74, 310–319, 2000. Coles, S.K. and Dick, T.E., Neurones in the ventrolateral pons are required for post-hypoxic frequency decline in rats, J. Physiol. 497, 79–94, 1996. Bach, K.B., Kinkead, R. and Mitchell, G.S., Post-hypoxia frequency decline in rats: Sensitivity to repeated hypoxia and a2-adrenoreceptor antagonism, Brain Res. 817, 25–33, 1999. Zabka, A.G., Behan, M. and Mitchell, G.S., Long-term facilitation of respiratory motor output decreases with age in male rats, J. Physiol. 531, 509–514, 2001. Zabka, A.G., Behan, M. and Mitchell, G.S., Time-dependent hypoxic respiratory responses in female rats are influenced by age and by the estrus cycle, J. Appl. Physiol. 91, 2831–2838, 2001. Coles, S.K., Miller, R., Huela, J., Wolken, P. and Schlenker, E., Frequency responses to hypoxia and hypercapnia in carotid body-denervated conscious rats, Respir. Physiol. Neurobiol. 130, 113–120, 2002. Guyenet, P.G., Koshiya, N., Huangfu, D., Verberne, A.J. and Riley, T.A., Central respiratory control of A5 and A6 pontine noradrenergic neurons, Am. J. Physiol. 264, R1035–R1044, 1993. Hedrick, M.S., Ryan, M.L., Pizarro, J. and Bisgard, G.E., Modulation of respiratory rhythm by alpha 2-adrenoceptors in awake and anesthetized goats, J. Appl. Physiol. 77, 742–750, 1994.
Respiratory Neuroplasticity 88.
89.
90.
91.
92.
93.
94.
95. 96.
97.
98.
99.
100. 101.
102.
103.
205
Coles, S.K., Ernsberger, P. and Dick, T.E., Post-hypoxic frequency decline does not depend on a2-adrenergic receptors in the adult rat, Brain Res. 794, 267–273, 1998. Clark, F.M. and Proudfit, H.K., Anatomical evidence for genetic differences in the innervation of the rat spinal cord by noradrenergic locus coeruleus neurons, Brain Res. 591, 44–53, 1992. Fuller, D.D., Baker, T.L., Behan, M. and Mitchell, G.S., Expression of hypoglossal long-term facilitation differs between substrains of Sprague– Dawley rat, Physiol. Genomics 4, 175–181, 2001. Kinkead, R. and Mitchell, G.S., Time-dependent hypoxic ventilatory responses in rats: Effects of ketanserin and 5-carboxamidotryptamine, Am. J. Physiol. 277, R658–R666, 1999. Coles, S.K., Ernsberger, P. and Dick, T.E., A role for NMDA receptors in posthypoxic frequency decline in the rat, Am. J. Physiol. 274, R1546–R1555, 1998. Ling, L., Fuller, D.D., Bach, K.B., Kinkead, R., Olson, E.B., Jr. and Mitchell, G.S., Chronic intermittent hypoxia elicits serotonin-dependent plasticity in the central neural control of breathing, J. Neurosci. 21, 5381– 5388, 2001. Baker, T.L. and Mitchell, G.S., Episodic but not continuous hypoxia elicits long-term facilitation of phrenic motor output in rats, J. Physiol. 529, 215–219, 2000. Fuller, D.D., Bach, K.B., Baker, T.L., Kinkead, R. and Mitchell, G.S., Long term facilitation of phrenic motor output, Respir. Physiol. 121, 135–146, 2000. Turner, D.L. and Mitchell, G.S., Long-term facilitation of ventilation following repeated hypoxic episodes in awake goats, J. Physiol. 499, 543– 550, 1997. Olson, E.B., Jr., Bohne, C.J., Dwinell, M.R., Podolsky, A., Vidruk, E.H., Fuller, D.D., Powell, F.L. and Mitchell, G.S., Ventilatory long-term facilitation in unanesthetized rats, J. Appl. Physiol. 91, 709–716, 2001. Dwinell, M.R., Janssen, P.L. and Bisgard, G.E., Lack of long-term facilitation of ventilation after exposure to hypoxia in goats, Respir. Physiol. 108, 1–9, 1997. Peng, Y.J. and Prabhakar, N.R., Reactive oxygen species in the plasticity of respiratory behavior elicited by chronic intermittent hypoxia, J. Appl. Physiol. 94, 2342–2349, 2003. Babcock, M.A. and Badr, M.S., Long-term facilitation of ventilation in humans during NREM sleep, Sleep 21, 709–716, 1998. Shkoukani, M., Babcock, M.A. and Badr, M.S., Effect of episodic hypoxia on upper airway mechanics in humans during NREM sleep, J. Appl. Physiol. 92, 2565–2570, 2002. Babcock, M.A., Shkoukani, M., Aboubakr, S.E. and Badr, M.S., Determinants of long-term facilitation in humans during NREM sleep, J. Appl. Physiol. 94, 53–59, 2003. Cao, K.Y., Zwillich, C.W., Berthon-Jones, M. and Sullivan, C.E., Increased normoxic ventilation induced by repetitive hypoxia in conscious dogs, J. Appl. Physiol. 73, 2083–2088, 1992.
206
Fuller et al.
104. McGuire, M., Zhang, Y., White, D.P. and Ling, L., Effect of hypoxic episode number and severity on ventilatory long-term facilitation in awake rats, J. Appl. Physiol. 93, 2155–2161, 2002. 105. Janssen, P.L. and Fregosi, R.F., No evidence for long-term facilitation after episodic hypoxia in spontaneously breathing, anesthetized rats, J. Appl. Physiol. 89, 1345–1351, 2000. 106. Jordan, A.S., Catcheside, P.G., O’Donoghue, F.J. and McEvoy, R.D., Longterm facilitation of ventilation is not present during wakefulness in healthy men or women, J. Appl. Physiol. 93, 2129–2136, 2002. 107. Peng, Y., Kline, D.D., Dick, T.E. and Prabhakar, N.R., Chronic intermittent hypoxia enhances carotid body chemoreceptor response to low oxygen, Adv. Exp. Med. Biol. 499, 33–38, 2001. 108. Bavis, R.W. and Mitchell, G.S., Intermittent hypoxia induces phrenic longterm facilitation in carotid denervated rats, J. Appl. Physiol. 94, 399–409, 2003. 109. Behan, M., Zabka, A.G. and Mitchell, G.S., Age and gender effects on serotonin-dependent plasticity in respiratory motor control, Respir. Physiol. Neurobiol. 131, 65–77, 2002. 110. Behan, M., Zabka, A.G., Thomas, C.F. and Mitchell, G.S., Sex steroid hormones and the neural control of breathing, Respir. Physiol. Neurobiol. 136, 249–263, 2003. 111. Bavis, R.W., Baker-Herman, T.L., Zabka, A.G., Golder, F.J., Fuller, D.D., Behan, M. and Mitchell, G.S., Respiratory long-term facilitation differs among inbred rat strains (abstr), FASEB J. 17, A824, 2003. 112. Mateika, J.H. and Fregosi, R.F., Long-term facilitation of upper airway muscle activities in vagotomized and vagally intact cats, J. Appl. Physiol. 82, 419–425, 1997. 113. Aboubakr, S.E., Taylor, A., Ford, R., Siddiqi, S. and Badr, M.S., Long-term facilitation in obstructive sleep apnea patients during NREM sleep, J. Appl. Physiol. 91, 2751–2757, 2001. 114. Millhorn, D.E., Eldridge, F.L. and Waldrop, T.G., Prolonged stimulation of respiration by endogenous central serotonin, Respir. Physiol. 42, 171–188, 1980. 115. Fregosi, R.F. and Mitchell, G.S., Long-term facilitation of inspiratory intercostal nerve activity following carotid sinus nerve stimulation in cats, J. Physiol. 477, 469–479, 1994. 116. Bach, K.B. and Mitchell, G.S., Hypoxia-induced long-term facilitation of respiratory activity is serotonin dependent, Respir. Physiol. 104, 251–260, 1996. 117. Fuller, D.D., Zabka, A.G., Baker, T.L. and Mitchell, G.S., Phrenic long-term facilitation requires 5-HT receptor activation during but not following episodic hypoxia, J. Appl. Physiol. 90, 2001–2006, 2001. 118. McCrimmon, D.R., Dekin, M.S. and Mitchell, G.S., Glutamate, GABA, and serotonin in ventilatory control, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 151–218, 1995. 119. Di Pasquale, E., Lindsay, A., Feldman, J., Monteau, R. and Hilaire, G., Serotonergic inhibition of phrenic motoneuron activity: An in vitro study in neonatal rat, Neurosci. Lett. 230, 29–32, 1997.
Respiratory Neuroplasticity
207
120. Baker-Herman, T.L. and Mitchell, G.S., Phrenic long-term facilitation requires spinal serotonin receptor activation and protein synthesis, J. Neurosci. 22, 6239–6246, 2002. 120.a. Backer-Herman, T.L., Fuller, D.D., Bavis, R.W., Zabka, A.G., Golder, F.J., Doperalski, N.J., Johnson, R.A., Watters, J.J. and Mitchell, G.S., BDNF is necessary and sufficient for spinal respiratory plasticity following intermittent hypoxia, Nat. Neurosci. 7, 48–55, 2004. 121. Gozal, D. and Gozal, E., Episodic hypoxia enhances late hypoxic ventilation in developing rat: Putative role of neuronal NO synthase, Am. J. Physiol. 276, R17–R22, 1999. 122. Tian, R.X., Kimura, S., Kondou, N., Fujisawa, Y., Zhou, M.S., Yoneyama, H., Kosaka, H., Rahman, M., Nishiyama, A. and Abe, Y., DOI, a 5-HT2 receptor agonist, induces renal vasodilation via nitric oxide in anesthetized dogs, Eur. J. Pharmacol. 437, 79–84, 2002. 123. Nozik-Grayck, E., McMahon, T.J., Huang, Y.C., Dieterle, C.S., Stamler, J.S. and Piantadosi, C.A., Pulmonary vasoconstriction by serotonin is inhibited by S-nitrosoglutathione, Am. J. Physiol. 282, L1057–L1065, 2002. 124. Millhorn, D.E., Eldridge, F.L. and Waldrop, T.G., Prolonged stimulation of respiration by a new central neural mechanism, Respir. Physiol. 41, 87–103, 1980. 125. Morris, K.F., Baekey, D.M., Shannon, R. and Lindsey, B.G., Respiratory neural activity during long-term facilitation, Respir. Physiol. 121, 119–133, 2000. 126. Morris, K.F., Shannon, R. and Lindsey, B.G., Changes in cat medullary neurone firing rates and synchrony following induction of respiratory longterm facilitation, J. Physiol. 532, 483–497, 2001. 127. Fuller, D.D., Johnson, S.M. and Mitchell, G.S., Respiratory long-term facilitation is associated with enhanced spinally-evoked phrenic potentials (abstr), Abstracts Viewer. Society for Neuroscience CD-ROM, 363.1, 2002. 128. Kinkead, R., Zhan, W.Z., Prakash, Y.S., Bach, K.B., Sieck, G.C. and Mitchell, G.S., Cervical dorsal rhizotomy enhances serotonergic innervation of phrenic motoneurons and serotonin-dependent long-term facilitation of phrenic motor output in rats, J. Neurosci. 18, 8436–8443, 1998. 129. Erickson, J.T. and Millhorn, D.E., Hypoxia and electrical stimulation of the carotid sinus nerve induce Fos-like immunoreactivity within catecholaminergic and serotoninergic neurons of the rat brainstem, J. Comp. Neurol. 348, 161–182, 1994. 130. Teppema, L.J., Veening, J.G., Kranenburg, A., Dahan, A., Berkenbosch, A. and Olievier, C., Expression of c-fos in the rat brainstem after exposure to hypoxia and to normoxic and hyperoxic hypercapnia, J. Comp. Neurol. 388, 169–190, 1997. 131. Basura, G.J, Zhou, S.Y., Walker, P.D. and Goshgarian, H.G., Distribution of serotonin 2A and 2C receptor mRNA expression in the cervical ventral horn and phrenic motoneurons following spinal cord hemisection, Exp. Neurol. 169, 255–263, 2001.
208
Fuller et al.
132. Chitravanshi, V.C. and Sapru, H.N., NMDA as well as non-NMDA receptors mediate the neurotransmission of inspiratory drive to phrenic motoneurons in the adult rat, Brain Res. 715, 104–112, 1996. 133. Chitravanshi, V.C. and Sapru, H.N., NMDA as well as non-NMDA receptors in phrenic nucleus mediate respiratory effects of carotid chemoreflex, Am. J. Physiol. 272, R302–R310, 1997. 134. Slack, S.E. and Thompson, S.W., Brain-derived neurotrophic factor induces NMDA receptor 1 phosphorylation in rat spinal cord, Neuroreport 13, 1967–1970, 2002. 135. Malinow, R. and Malenka, R.C., AMPA receptor trafficking and synaptic plasticity, Annu. Rev. Neurosci. 25, 103–126, 2002. 136. Schinder, A.F. and Poo, M., The neurotrophin hypothesis for synaptic plasticity, Trends Neurosci. 23, 639–645, 2000. 137. Dempsey, J.A. and Forster, H.V., Mediation of ventilatory adaptations, Physiol. Rev. 62, 262–346, 1982. 138. Powell, F.L., Huey, K.A. and Dwinell, M.R., Central nervous system mechanisms of ventilatory acclimatization to hypoxia, Respir. Physiol. 121, 223–236, 2000. 139. Moore, L.G., Comparative human ventilatory adaptation to high altitude, Respir. Physiol. 121, 257–276, 2000. 140. Busch, M.A., Bisgard, G.E. and Forster, H.V., Ventilatory acclimatization to hypoxia is not dependent on arterial hypoxemia, J. Appl. Physiol. 58, 1874–1880, 1985. 141. Engwall, M.J., Vidruk, E.H., Nielsen, A.M. and Bisgard, G.E., Response of the goat carotid body to acute and prolonged hypercapnia, Respir. Physiol. 74, 335–344, 1988. 142. Nielsen, A.M., Bisgard, G.E. and Vidruk, E.H., Carotid chemoreceptor activity during acute and sustained hypoxia in goats, J. Appl. Physiol. 65, 1796–1802, 1988. 143. Vizek, M., Pickett, C.K. and Weil, J.V., Increased carotid body hypoxic sensitivity during acclimatization to hypobaric hypoxia, J. Appl. Physiol. 63, 2403–2410, 1987. 144. He, L., Chen, J., Dinger, B., Stensaas, L. and Fidone, S., Endothelin modulates chemoreceptor cell function in mammalian carotid body, Adv. Exp. Med. Biol. 410, 305–311, 1996. 145. Chen, J., He, L., Dinger, B. and Fidone, S., Pharmacological effects of endothelin in rat carotid body. Activation of second messenger pathways and potentiation of chemoreceptor activity, Adv. Exp. Med. Biol. 475, 517–525, 2000. 146. Chen, J., He, L., Dinger, B., Stensaas, L. and Fidone, S., Role of endothelin and endothelin A-type receptor in adaptation of the carotid body to chronic hypoxia, Am. J. Physiol. 282, L1314–L1323, 2002. 146a. Wang, Z.Y. and Bisgard, G.E., Chronic hypoxia-induced morphological and neurochemical changes in the carotid body, Microsc. Res. Tech. 59, 168–177, 2002.
Respiratory Neuroplasticity
209
147. Dwinell, M.R. and Powell, F.L., Chronic hypoxia enhances the phrenic nerve response to arterial chemoreceptor stimulation in anesthetized rats, J. Appl. Physiol. 87, 817–823, 1999. 148. Huey, K.A. and Powell, F.L., Time-dependent changes in dopamine D(2)receptor mRNA in the arterial chemoreflex pathway with chronic hypoxia, Brain Res. Mol. Brain Res. 75, 264–270, 2000. 149. Huey, K.A., Brown, I.P., Jordan, M.C. and Powell, F.L., Changes in dopamine D(2)-receptor modulation of the hypoxic ventilatory response with chronic hypoxia, Respir. Physiol. 123, 177–187, 2000. 150. Soulier, V., Gestreau, C., Borghini, N., Dalmaz, Y., Cottet-Emard, J.M. and Pequignot, J.M., Peripheral chemosensitivity and central integration: Neuroplasticity of catecholaminergic cells under hypoxia, Comp. Biochem. Physiol. A 118, 1–7, 1997. 151. Dwinell, M.R,, Huey, K.A. and Powell, F.L., Chronic hypoxia induces changes in the central nervous system processing of arterial chemoreceptor input, Adv. Exp. Med. Biol. 475, 477–484, 2000. 152. Serebrovskaya, T.V., Intermittent hypoxia research in the former Soviet Union and the Commonwealth of Independent States: History and review of the concept and selected applications, High Alt. Med. Biol. 3, 205–221, 2002. 153. Wilber, R.L., Current trends in altitude training, Sports Med. 31, 249–265, 2001. 154. Fletcher, E.C., Physiological consequences of intermittent hypoxia: Systemic blood pressure, J. Appl. Physiol. 90, 1600–1605, 2001. 155. Greenberg, H.E., Sica, A., Batson, D. and Scharf, S.M., Chronic intermittent hypoxia increases sympathetic responsiveness to hypoxia and hypercapnia, J. Appl. Physiol. 86, 298–305, 1999. 156. Gozal, E., Row, B.W., Schurr, A. and Gozal, D., Developmental differences in cortical and hippocampal vulnerability to intermittent hypoxia in the rat, Neurosci. Lett. 305, 197–201, 2001. 157. Gozal, E., Gozal, D., Pierce, W.M., Thongboonkerd, V., Scherzer, J.A., Sachleben, L.R., Jr., Brittian, K.R., Guo, S.Z., Cai, J. and Klein, J.B., Proteomic analysis of CA1 and CA3 regions of rat hippocampus and differential susceptibility to intermittent hypoxia, J. Neurochem. 83, 331–345, 2002. 158. Gozal, D. and Gozal, E., Respiratory plasticity following intermittent hypoxia: Developmental interactions, J. Appl. Physiol. 90, 1995–1999, 2001. 159. Berger, K.I., Ayappa, I., Sorkin, I.B., Norman, R.G., Rapoport, D.M. and Goldring, R.M., Postevent ventilation as a function of CO2 load during respiratory events in obstructive sleep apnea, J. Appl. Physiol. 93, 917–924, 2002. 160. Narkiewicz, K., van de Borne, P.J., Pesek, C.A., Dyken, M.E., Montano, N. and Somers, V.K., Selective potentiation of peripheral chemoreflex sensitivity in obstructive sleep apnea, Circulation 99, 1183–1189, 1999. 161. Benlloch, E., Cordero, P., Morales, P., Soler, J.J. and Macian, V., Ventilatory pattern at rest and response to hypercapnic stimulation in patients with obstructive sleep apnea syndrome, Respiration 62, 4–9, 1995.
210
Fuller et al.
162. Sin, D.D., Jones, R.L. and Man, G.C., Hypercapnic ventilatory response in patients with and without obstructive sleep apnea: Do age, gender, obesity, and daytime PaCO2 matter? Chest 117, 454–459, 2000. 163. Sullivan, C.E., Grunstein, R.R., Maronne, O. and Berthon-Jones, M., Sleep apnea pathophysiology: Upper airway and control of breathing, in Obstructive Sleep Apnea Syndrome: Clinical Research and Treatments, Guilleminault, C. and Partinen, M., eds., New York, Raven Press, pp. 49–69, 1990. 164. Fuller, D.D., Wang, Z.Y., Ling, L., Olson, E.B., Bisgard, G.E. and Mitchell, G.S., Induced recovery of hypoxic phrenic responses in adult rats exposed to hyperoxia for the first month of life, J. Physiol. 536, 917–926, 2001. 165. Serebrovskaya, T.V., Karaban, I.N., Kolesnikova, E.E., Mishunina, T.M., Kuzminskaya, L.A., Serbrovsky, A.N. and Swanson, R.J., Human hypoxic ventilatory response with blood dopamine content under intermittent hypoxic training, Can. J. Physiol. Pharmacol. 77, 967–973, 1999. 166. Bernardi, L., Passino, C., Serebrovskaya, Z., Serebrovskaya, T. and Appenzeller, O., Respiratory and cardiovascular adaptations to progressive hypoxia; Effect of interval hypoxic training, Eur. Heart J. 22, 879–886, 2001. 167. Mahamed, S. and Duffin, J., Repeated hypoxic exposures change respiratory chemoreflex control in humans, J. Physiol. 534, 595–603, 2001. 168. Katayama, K., Shima, N., Sato, Y., Qiu, J.C., Ishida, K., Mori, S. and Miyamura, M., Effect of intermittent hypoxia on cardiovascular adaptations and response to progressive hypoxia in humans, High Alt. Med. Biol. 2, 501–508, 2001. 169. Ricart, A., Casas, H., Casas, M., Pages, T., Palacios, L., Rama, R., Rodriguez, F.A., Viscor, G. and Ventura, J.L., Acclimatization near home? Early respiratory changes after short-term intermittent exposure to simulated altitude, Wilderness Environ. Med. 11, 84–88, 2000. 170. Garcia. N., Hopkins, S.R. and Powell, F.L., Effects of intermittent hypoxia on the isocapnic hypoxic ventilatory response and erythropoiesis in humans, Respir. Physiol. 123, 39–49, 2000. 171. Richalet, J.P., Donoso, M.V., Jimenez, D., Antezana, A.M., Hudson, C., Cortes, G., Osorio, J. and Leon, A., Chilean miners commuting from sea level to 4500 m: A prospective study, High Alt. Med. Biol. 3, 159–166, 2002. 172. Townsend, N.E., Gore, C.J., Hahn, A.G., McKenna, M.J., Aughey, R.J., Clark, S.A., Kinsman, T., Hawley, J.A. and Chow, C.M., Living hightraining low increases hypoxic ventilatory response of well-trained endurance athletes, J. Appl. Physiol. 93, 1498–1505, 2002. 173. Honda, Y., Tani, H., Masuda, A., Kobayashi, T., Nishino, T., Kimura, H., Masuyama, S. and Kuriyama, T., Effect of prior O2 breathing on ventilatory response to sustained isocapnic hypoxia in adult humans, J. Appl. Physiol. 81, 1627–1632, 1996. 174. Gozal, D., Potentiation of hypoxic ventilatory response by hyperoxia in the conscious rat: Putative role of nitric oxide, J. Appl. Physiol. 85, 129–132, 1998. 175. Lahiri, S., Mulligan, E., Andronikou, S., Shirahata, M. and Mokashi, A., Carotid body chemosensory function in prolonged normobaric hyperoxia in the cat, J. Appl. Physiol. 62, 1924–1931, 1987.
Respiratory Neuroplasticity
211
176. Liberzon, I., Arieli, R. and Kerem, D., Attenuation of hypoxic ventilation by hyperbaric O2: Effects of pressure and exposure time, J. Appl. Physiol. 66, 851–856, 1989. 177. Lahiri, S., Mokashi, A., Shirahata, M. and Andronikou, S., Chemical respiratory control in chronically hyperoxic cats, Respir. Physiol. 82, 201–215, 1990. 178. Mokashi, A., Lahiri, S., Aortic and carotid body chemoreception in prolonged hyperoxia in the cat, Respir. Physiol. 86, 233–243, 1991. 179. Mokashi, A., Di Guilio, C., Morelli, L. and Lahiri, S., Chronic hyperoxic effects on cat carotid body catecholamines and structure, Respir. Physiol. 97, 25–32, 1994. 180. Di Giulio, C., Di Muzio, M., Sabatino, G., Spoletini, L., Amicarelli, F., Di Ilio, C. and Modesti, A., Effect of chronic hyperoxia on young and old rat carotid body ultrastructure, Exp. Gerontol. 33, 319–329, 1998. 181. Ren, X., Fatemian, M. and Robbins, P.A., Changes in respiratory control in humans induced by 8 h of hyperoxia, J. Appl. Physiol. 89, 655–662, 2000. 182. Gelfand, R., Lambertsen, C.J., Clark, J.M. and Hopkin, E., Hypoxic ventilatory sensitivity in men is not reduced by prolonged hyperoxia (Predictive Studies V and VI), J. Appl. Physiol. 84, 292–302, 1998. 183. Birchard, G.F., Kilgore, D.L., Jr. and Boggs, D.F., Respiratory gas concentrations and temperatures within the burrows of three species of burrow-nesting birds, Wilson Bull. 96, 451–456, 1984. 184. Tenney, S.M. and Boggs, D.F., Comparative mammalian respiratory control, in Handbook of Physiology: Control of Breathing, Cherniack, N.S. and Widdicombe, J.G., eds., Bethesda, American Physiological Society, pp. 833–855, 1986. 185. Feldman, J.L., Neurophysiology of breathing in mammals, in Handbook of Physiology: Control of Breathing, Cherniack, N.S. and Widdicombe, J.D., eds., Bethesda, American Physiological Society, pp. 463–524, 1986. 186. Dong, X.W., Gozal, D., Rector, D.M. and Harper, R.M., Ventilatory CO2induced optical activity changes of cat ventral medullary surface, Am. J. Physiol. 265, R494–R503, 1993. 187. Bach, K.B. and Mitchell, G.S., Hypercapnia-induced long-term depression of respiratory activity requires a2-adrenergic receptors, J. Appl. Physiol. 84, 2099–2105, 1998. 188. Gozal, D., Ben-Ari, J.H., Harper, R.M. and Keens, T.G., Ventilatory responses to repeated short hypercapnic challenges, J. Appl. Physiol. 78, 1374–1381, 1995. 189. Gozal, D., Arens, R., Omlin, K.J., Ben-Ari, J.H., Aljadeff, G., Harper, R.M. and Keens, T.G., Ventilatory response to consecutive short hypercapnic challenges in children with obstructive sleep apnea, J. Appl. Physiol. 79, 1608–1614, 1995. 190. Schaefer, K.E., Hastings, B.J., Carey, C.R. and Nichols, G., Jr., Respiratory acclimatization to carbon dioxide, J. Appl. Physiol. 18, 1071–1078, 1963. 191. Lai, Y.L., Lamm, J.E. and Hildebrandt, J., Ventilation during prolonged hypercapnia in the rat, J. Appl. Physiol. 51, 78–83, 1981.
212
Fuller et al.
192. Kondo, T., Kumagai, M., Ohta, Y. and Bishop, B., Ventilatory responses to hypercapnia and hypoxia following chronic hypercapnia in the rat, Respir. Physiol. 122, 35–43, 2000. 193. Seifert, E.L. and Mortola, J.P., Circadian pattern of ventilation during acute and chronic hypercapnia in conscious adult rats, Am. J. Physiol. 282, R244–R251, 2002. 194. Bebout, D.E. and Hempleman, S.C., Chronic hypercapnia resets CO2 sensitivity of avian intrapulmonary chemoreceptors, Am. J. Physiol. 276, R317–R322, 1999. 195. Scalise, R., Bavis, R.W. and Kilgore, D.L. Jr., Hypercapnic ventilatory response of Japanese quail before and after chronic exposure to moderate CO2 (abstr), FASEB J. 14, A79, 2000. 196. Berardi, N., Pizzorusso, T. and Maffei, L., Critical periods during sensory development, Curr. Opin. Neurobiol. 10, 138–145, 2000. 197. Gleed, R.D. and Mortola, J.P., Ventilation in newborn rats after gestation at simulated high altitude, J. Appl. Physiol. 70, 1146–1151, 1991. 198. Olson, E.B., Jr. and Dempsey, J.A., Rat as a model for humanlike ventilatory adaptation to chronic hypoxia, J. Appl. Physiol. 44, 763–769, 1978. 199. Peyronnet, J., Roux, J.C., Ge´loe¨n, A., Tang, L.Q., Pequignot, J.M., Lagercrantz, H. and Dalmaz, Y., Prenatal hypoxia impairs the postnatal development of neural and functional chemoafferent pathway in rat, J. Physiol. (Lond) 524, 525–537, 2000. 200. Cheung, E., Wong, N. and Mortola, J.P., Compliance of the respiratory system in newborn and adult rats after gestation in hypoxia, J. Comp. Physiol. B 170, 193–199, 2000. 201. White, L.D. and Lawson, E.E., Effects of chronic prenatal hypoxia on tyrosine hydroxylase and phenylethanolamine N-methyltransferase messanger RNA and protein levels in medulla oblongata of postnatal rat, Pediatr. Res. 42, 455–462, 1997. 202. Hempleman, S.C., Sodium and potassium current in neonatal rat carotid body cells following chronic in vivo hypoxia, Brain Res. 699, 42–50, 1995. 203. Hempleman, S.C., Increased calcium current in carotid body glomus cells following in vivo acclimatization to chronic hypoxia, J. Neurophysiol. 76, 1880–1886, 1996. 204. Moss, T.J., Davey, M.G., McCrabb, G.J. and Harding, R., Development of ventilatory responsiveness to progressive hypoxia and hypercapnia in lowbirth-weight lambs, J. Appl. Physiol. 81, 1555–1561, 1996. 205. Harding, R., Tester, M.I., Moss, T.J., Davey, M.G., Louey, S., Joyce, B., Hooper, S.B. and Maritz, G., Effects of intra-uterine growth restriction on the control of breathing and lung development after birth, Clin. Exp. Pharmacol. Physiol. 27, 114–119, 2000. 206. Moss, T.J. and Harding, R., Ventilatory and arousal responses of sleeping lambs to respiratory challenges: Effect of prenatal maternal anemia, J. Appl. Physiol. 88, 641–648, 2000. 207. Moss, T.J. and Harding, R., Ventilatory and arousal responses to respiratory stimuli of full term, intrauterine growth restricted lambs, Respir. Physiol. 124, 195–204, 2001.
Respiratory Neuroplasticity
213
208. Tolcos, M., Rees, S., McGregor, H. and Walker, D., Consequences of intrauterine growth restriction on ventilatory and thermoregulatory responses to asphyxia and hypercapnia in the newborn guinea-pig, Reprod. Fertil. Dev. 14, 85–92, 2002. 209. McGregor, H.P., Westcott, K. and Walker, D.W., The effect of prenatal exposure to carbon monoxide on breathing and growth of the newborn guinea pig, Pediatr. Res. 43, 126–131, 1998. 210. Mortola, J.P., Morgan, C.A. and Virgona, V., Respiratory adaptation to chronic hypoxia in newborn rats, J. Appl. Physiol. 61, 1329–1336, 1986. 211. Dotta, A. and Mortola, J.P., Postnatal development of the denervated lung in normoxia, hypoxia, or hyperoxia, J. Appl. Physiol. 73, 1461–1466, 1992. 212. Okubo, S. and Mortola, J.P., Long-term respiratory effects of neonatal hypoxia in the rat, J. Appl. Physiol. 64, 952–958, 1988. 213. Okubo, S. and Mortola, J.P., Control of ventilation in adult rats hypoxic in the neonatal period, Am. J. Physiol. 259, R836–R841, 1990. 214. Sørensen, S.C. and Severinghaus, J.W., Respiratory insensitivity to acute hypoxia persisting after correction of tetralogy of Fallot, J. Appl. Physiol. 25, 221–223, 1968. 215. Edelman, N.H., Lahiri, S., Braudo, M.D., Cherniack, N.S. and Fishman, A.P., The blunted ventilatory response to hypoxia in cyanotic congenital heart disease, N. Engl. J. Med. 282, 405–411, 1970. 216. Blesa, M.I., Lahiri, S., Rashkind, W.J. and Fishman, A.P., Normalization of the blunted ventilatory response to acute hypoxia in congenital cyanotic heart disease, N. Engl. J. Med. 296, 237–241, 1977. 217. Gamboa, A., Le´on-Velarde, F., Rivera-Ch, M., Palacios, J.-A., Pragnell, T.R., O’Connor, D.F. and Robbins, P.A., Acute and sustained ventilatory responses to hypoxia in high-altitude natives living at sea level, J. Appl. Physiol. 94, 1255–1262, 2003. 218. Sørensen, S.C. and Severinghaus, J.W., Irreversible respiratory insensitivity to acute hypoxia in man born at high altitude, J. Appl. Physiol. 25, 217–220, 1968. 219. Byrne-Quinn, E., Sodal, I.E. and Weil, J.V., Hypoxic and hypercapnic ventilatory drives in children native to high altitude, J. Appl. Physiol. 32, 44–46, 1972. 220. Lahiri, S., DeLaney, R.G., Brody, J.S., Simpser, M., Velasquez, T., Motoyama, E.K. and Polgar, C., Relative role of environmental and genetic factors in respiratory adaptation to high altitude, Nature 261, 133–135, 1976. 221. Lahiri, S., Adaptive respiratory regulation—lessons from high altitudes, in Environmental Physiology: Aging, Heat and Altitude, Hovarth, S.M. and Yousef, M.K., eds., New York, Elsevier North Holland, pp. 341–350, 1981. 222. Vargas, M., Leo´n-Velarde, F., Monge, C., Palacios, J.-A. and Robbins, P.A., Similar hypoxic ventilatory responses in sea-level natives and high-altitude natives living at sea level, J. Appl. Physiol. 84, 1024–1029, 1998. 223. Lahiri, S., Brody, J.S., Motoyama, E.K. and Velasquez, T.M., Regulation of breathing in newborns at high altitude, J. Appl. Physiol. 44, 673–678, 1978.
214
Fuller et al.
224. Weil, J.V., Byrne-Quinn, E., Sodal, I.E., Filley, G.F. and Grover, R.F., Acquired attenuation of chemoreceptor function in chronically hypoxic man at high altitude, J. Clin. Investig. 50, 186–194, 1971. 225. Fatemian, M., Gamboa, A., Le´on-Velarde, F., Rivera-Ch, M., Palacios, J.-A. and Robbins, P.A., Ventilatory response to CO2 in high-altitude natives and patients with chronic mountain sickness, J. Appl. Physiol. 94, 1279–1287, 2003. 226. Eden, G.J. and Hanson, M.A., Effects of chronic hypoxia from birth on the ventilatory response to acute hypoxia in the newborn rat, J. Physiol. (Lond.) 392, 11–19, 1987. 227. Hanson, M.A., Kumar, P. and Williams, B.A., The effect of chronic hypoxia upon the development of respiratory chemoreflexes in the newborn kitten, J. Physiol. (Lond.) 411, 563–574, 1989. 228. Hanson, M.A., Eden, G.J., Nijhuis, J.G. and Moore, P.J., Peripheral chemoreceptors and other oxygen sensors in the fetus and newborn, in Chemoreceptors and Reflexes in Breathing: Cellular and Molecular Aspects, Lahiri, S., Forster, R.E., Davies, R.O. and Pack, A.I., eds., New York, Oxford University Press, pp. 113–120, 1989. 229. Sladek, M., Parker, R.A., Grogaard, J.B. and Sundell, H.W., Long-lasting effect of prolonged hypoxemia after birth on the immediate ventilatory response to changes in arterial partial pressure of oxygen in young lambs, Pediatr. Res. 34, 821–828, 1993. 230. Wyatt, C.N., Wright, C., Bee, D. and Peers, C., O2-sensitive Kþ currents in carotid body chemoreceptor cells from normoxic and chronically hypoxic rats and their roles in hypoxic chemotransduction, Proc. Natl. Acad. Sci. USA 92, 295–299, 1995. 231. Mortola, J.P. and Saiki, C., Ventilatory response to hypoxia in rats: Gender differences, Respir. Physiol. 106, 21–34, 1996. 232. Bavis, R.W., Olson, E.B., Jr., Vidruk, E.H., Fuller, D.D. and Mitchell, G.S., Developmental plasticity of the hypoxic ventilatory response in rats induced by neonatal hypoxia, J. Physiol. 557, 645–660, 2004. 234. Hanson, M. and Kumar, P., Chemoreceptor function in the fetus and neonate, Adv. Exp. Med. Biol. 360, 99–108, 1994. 235. Hanson, M.A., Role of chemoreceptors in effects of chronic hypoxia, Comp. Biochem. Physiol. 119, 695–703, 1998. 236. Sterni, L.M., Bamford, O.S., Wasicko, M.J. and Carroll, J.L., Chronic hypoxia abolished the postnatal increase in carotid body type I cell sensitivity to hypoxia, Am. J. Physiol. 277, L645–L652, 1999. 237. Landauer, R.C., Pepper, D.R. and Kumar, P., Effect of chronic hypoxaemia from birth upon chemosensitivity in the adult rat carotid body in vitro, J. Physiol. (Lond.) 485, 543–550, 1995. 238. Okubo, S. and Mortola, J.P., Respiratory mechanics in adult rats hypoxic in the neonatal period, J. Appl. Physiol. 66, 1772–1778, 1989. 239. Kass, L.J. and Bazzy, A.R., Chronic hypoxia modulates diaphragm function in the developing rat, J. Appl. Physiol. 90, 2325–2329, 2001. 240. Gozal, D., Gozal, E., Reeves, S.R. and Lipton, A.J., Gasping and autoresuscitation in the developing rat: Effect of antecedent intermittent hypoxia, J. Appl. Physiol. 92, 1141–1144, 2002.
Respiratory Neuroplasticity
215
241. Matsuoka, T., Yoda, T., Ushikubo, S., Matsuzawa, S., Sasano, T. and Komiyama, A., Repeated acute hypoxia temporarily attenuates the ventilatory respiratory response to hypoxia in conscious newborn rats, Pediatr. Res. 46, 120–125, 1999. 242. Lowry, T.F., Rice, T.B., Forster, H.V. and Franciosi, R.A., The effect of perinatal episodic hypoxia (EH) on development of normal respiration and mean arterial pressure (MAP) in neonatal rats (abstr), FASEB J. 15, A817, 2001. 243. Saiki, C. and Mortola, J.P., Ventilatory control in infant rats after daily episode of anoxia, Pediatr. Res. 35, 490–493, 1994. 244. Saiki, C. and Matsumoto, S., Effect of neonatal anoxia on the ventilatory response to hypoxia in developing rats, Pediatr. Pulmonol. 28, 313–320, 1999. 245. Waters, K.A., Laferrie`re, A., Paquette, J., Goodyer, C. and Moss, I.R., Curtailed respiration by repeated vs. isolated hypoxia in maturing piglets is unrelated to NTS ME or SP levels, J. Appl. Physiol. 83, 522–529, 1997. 246. Waters, K.A., Beardsmore, C.S., Paquette, J., Meehan, B., Coˆte´, A. and Moss, I.R., Respiratory responses to rapid-onset, repetitive vs. continuous hypoxia in piglets, Respir. Physiol. 105, 135–142, 1996. 247. Waters, K.A. and Tinworth, K.D., Depression of ventilatory responses after daily, cyclic hypercapnic hypoxia in piglets, J. Appl. Physiol. 90, 1065–1073, 2001. 248. Ling, L., Olson, E.B., Jr., Vidruk, E.H. and Mitchell, G.S., Attenuation of the hypoxic ventilatory response in adult rats following one month of perinatal hyperoxia, J. Physiol. (Lond.) 495, 561–571, 1996. 249. Ling, L., Olson, E.B., Jr., Vidruk, E.H. and Mitchell, G.S., Phrenic responses to isocapnic hypoxia in adult rats following perinatal hyperoxia, Respir. Physiol. 109, 107–116, 1997. 249a. Bavis, R.W., Olson, E.B., Jr., Vidruk, E.H., Bisgard, G.E. and Mitchell, G.S., Level and duration of developmental hyperoxia influence impairment of hypoxic phrenic responses in rats, J. Appl. Physiol. 95, 1550–1559, 2003. 250. Eden, G.J. and Hanson, M.A., Effect of hyperoxia from birth on the carotid chemoreceptor and ventilatory responses of rats to acute hypoxia (abstr), J. Physiol. (Lond.) 374, 24P, 1986. 251. Fuller, D.D., Bavis, R.W., Vidruk, E.H., Wang, Z.Y., Olson, E.B., Bisgard, G.E. and Mitchell, G.S., Life-long impairment of hypoxic phrenic responses in rats following 1 month of developmental hyperoxia, J. Physiol. 538, 947–955, 2002. 252. Bavis, R.W., Olson, E.B., Jr. and Mitchell, G.S., Critical developmental period for hyperoxia-induced blunting of hypoxic phrenic responses in rats, J. Appl. Physiol. 92, 1013–1018, 2002. 253. Ling, L., Olson, E.B., Jr., Vidruk, E.H. and Mitchell, G.S., Integrated phrenic responses to carotid afferent stimulation in adult rats following perinatal hyperoxia, J. Physiol. (Lond.) 500, 787–796, 1997. 254. Erickson, J.T., Mayer, C., Jawa, A., Ling, L., Olson, E.B., Jr., Vidruk, E.H., Mitchell, G.S. and Katz, D.M., Chemoafferent degeneration and carotid body hypoplasia following chronic hyperoxia in newborn rats, J. Physiol. (Lond.) 509, 519–526, 1998.
216
Fuller et al.
255. Prieto-Lloret, J., Caceres, A.I., Obeso, A., Rigual, R., Rocher, A., Bustamante, R., Castan˜eda, J., Lo´pez-Lo´pez, J.R., Perez-Garcia, M.T., Agapito, A. and Gonza´lez, C., Effects of perinatal hyperoxia on carotid body chemoreceptor activity in vitro, in Chemoreception: From Cellular Signaling to Functional Plasticity, Pequignot, J.M., Gonza´lez, C., Nurse, C., Dalmaz, Y. and Prabhakar, N., eds., New York, Kluwer Academic/Plenum Publishers, 2003. 256. Bisgard, G.E., Mitchell, G.S., Wang, Z.Y. and Olson, E.B., Jr., Attenuated carotid body function after one week of postnatal hyperoxia in rats (abstr), FASEB J. 16, A68, 2002. 257. Carroll, J.L., Kim, I., Boyle, K.M., Carle, C.M. and Donnelly, D.F., Perinatal hyperoxia impairs responsiveness of single-unit, rat carotid chemoreceptor activity (abstr), FASEB J. 17, LB127, 2003. 258. Kim, I., Boyle, K.M., Carle, C.M., Donnelly, D.F. and Carroll, J.L., Perinatal hyperoxia reduces the depolarization and calcium increase in response to an acute hypoxia challenge in rat carotid chemoreceptor cells (abstr), FASEB J. 17, LB128, 2003. 259. Hertzberg, T., Brosenitsch, T. and Katz, D.M., Depolarizing stimuli induce high levels of dopamine synthesis in fetal rat sensory neurons, Neuroreport 7, 233–237, 1995. 260. Balkowiec, A. and Katz, D.M., Activity-dependent release of endogenous brain-derived neurotrophic factor from primary sensory neurons detected by ELISA in situ, J. Neurosci. 20, 7417–7423, 2000. 261. Hertzberg, T., Fan, G., Finley, J.C.W., Erickson, J.T. and Katz, D.M., BDNF supports mammalian chemoafferent neurons in vitro and following peripheral target removal in vivo, Dev. Biol. 166, 801–811, 1994. 262. Paulding, W.R., Schnell, P.O., Bauer, A.L., Striet, J.B., Nash, J.A., Kuznetsova, A.V. and Czyzyk-Krzeska, M.F., Regulation of gene expression for neurotransmitters during adaptation to hypoxia in oxygen-sensitive neuroendocrine cells, Microsc. Res. Tech. 59, 178–187, 2002. 263. D’Angio, C.T. and Finkelstein, J.N., Oxygen regulation of gene expression: A study in opposites, Mol. Genet. Metab. 71, 371–380, 2000. 264. Dor, Y., Porat, R. and Keshet, E., Vascular endothelial growth factor and vascular adjustments to perturbations in oxygen homeostasis, Am. J. Physiol. 280, C1367–C1374, 2001. 265. Jamieson, D., Chance, B., Cadenas, E. and Boveris, A., The relation of free radical production to hyperoxia, Annu. Rev. Physiol. 48, 703–719, 1986. 266. Frank, L., Developmental aspects of experimental pulmonary oxygen toxicity, Free Rad. Biol. Med. 11, 463–494, 1991. 267. Chandel, N.S. and Schumacker, P.T., Cellular oxygen sensing by mitochondria: Old questions, new insight, J. Appl. Physiol. 88, 1880–1889, 2000. 268. Prabhakar, N.R., Oxygen sensing during intermittent hypoxia: Cellular and molecular mechanisms, J. Appl. Physiol. 90, 1986–1994, 2001. 269. Dro¨ge, W., Free radicals in the physiological control of cell function, Physiol. Rev. 82, 47–95, 2002. 270. Claure, N., Gerhardt, T., Everett, R., Musante, G., Herrera, C. and Bancalari, E., Closed-loop controlled inspired oxygen concentration for mechanically
Respiratory Neuroplasticity
271.
272.
273.
274.
275.
276.
277. 278. 279.
280.
281. 282.
283.
284.
285.
217
ventilated very low birth weight infants with frequent episodes of hypoxemia, Pediatrics 107, 1120–1124, 2001. Katz-Salamon, M. and Lagercrantz, H., Hypoxic ventilatory defence in very preterm infants: Attenuation after long-term oxygen treatment, Arch. Dis. Child 70, F90–F95, 1994. Calder, N.A., Williams, B.A., Smyth, J., Boon, A.W., Kumar, P. and Hanson, M.A., Absence of ventilatory response to alternating breaths of mild hypoxia and air in infants who have had bronchopulmonary dysplasia: Implications for the risk of sudden infant death, Pediatr. Res. 35, 677–681, 1994. Katz-Salamon, M., Jonsson, B. and Lagercrantz, H., Blunted peripheral chemoreceptor response to hyperoxia in a group of infants with bronchopulmonary dysplasia, Pediatr. Pulmonol. 20, 101–106, 1995. Katz-Salamon, M., Eriksson, M. and Jonsson, B., Development of peripheral chemoreceptor function in infants with chronic lung disease and initially lacking hyperoxic response, Arch. Dis. Child 75, F4–F9, 1996. Varughese, M., Patole, S., Shama, A. and Whitehall, J., Permissive hypercapnia in neonates: The case of the good, the bad, and the ugly, Pediatr. Pulmonol. 33, 56–64, 2002. Birchard, G.F., Boggs, D.F. and Tenney, S.M., Effect of perinatal hypercapnia on the adult ventilatory response to carbon dioxide, Respir. Physiol. 57, 341–347, 1984. Rezzonico, R. and Mortola, J.P., Respiratory adaptation to hypercapnia in newborn rats and its long term effects (abstr), Physiologist 31, A171, 1988. Rezzonico, R. and Mortola, J.P., Respiratory adaptation to chronic hypercapnia in newborn rats, J. Appl. Physiol. 67, 311–315, 1989. Bavis, R.W. and Kilgore, D.L., Jr., Effects of embryonic CO2 exposure on the adult ventilatory response in quail: Does gender matter? Respir. Physiol. 126, 183–199, 2001. Williams, B.R., Jr. and Kilgore, D.L., Jr., Ontogenetic modification of the hypercapnic ventilatory response in the zebra finch, Respir. Physiol. 90, 125–134, 1992. Rezzonico, R., Gleed, R.D. and Mortola, J.P., Respiratory mechanics in adult rats hypercapnic in the neonatal period, J. Appl. Physiol. 68, 2274–2279, 1990. Mortola, J.P. and Gautier, H., Interaction between metabolism and ventilation: Effects of respiratory gases and temperature, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 1011–1064, 1995. Sant’Anna, G.M. and Mortola, J.P., Thermal and respiratory control in young rats with altered caloric intake during postnatal development, Respir. Physiol. Neurobiol. 133, 215–227, 2002. Sant’Anna, G.M. and Mortola, J.P., Thermal and respiratory control in young rats exposed to cold during postnatal development, Comp. Biochem. Physiol. A 134, 449–459, 2003. Lowry, T.F., Forster, H.V., Pan, L.G., Serra, A., Wenninger, J., Nash, R., Sheridan, D. and Franciosi, R.A., Effects on breathing of carotid body denervation in neonatal piglets, J. Appl. Physiol. 87, 2128–2135, 1999.
218
Fuller et al.
286. Bisgard, G.E., Forster, H.V. and Klein, J.P., Recovery of peripheral chemoreceptor function after denervation in ponies, J. Appl. Physiol. 49, 964–970, 1980. 287. Smith, P.G. and Mills, E., Restoration of reflex ventilatory response to hypoxia after removal of carotid bodies in the cat, Neuroscience 5, 573–580, 1980. 288. Majumdar, S., Smith, P.G. and Mills, E., Evidence for central reorganisation of ventilatory chemoreflex pathways in the cat during regeneration of visceral afferents in the carotid sinus nerve, Neuroscience 7, 1309–1316, 1982. 289. Martin-Body, R.L., Robson, G.J. and Sinclair, J.D., Restoration of hypoxic ventilatory responses in the awake rat after carotid body denervation by sinus nerve section, J. Physiol. (Lond.) 380, 61–73, 1986. 290. Roux, J.C., Peyronnet, J., Pascual, O., Dalmaz, Y. and Pequignot, J.M., Ventilatory and central neurochemical reorganization of O2 chemoreflex after carotid sinus nerve transection in rat, J. Physiol. (Lond.) 522, 493–501, 2000. 291. Roux, J.C., Pequignot, J.M., Dumas, S., Pascual, O., Ghilini, G., Pequignot, J., Mallet, J. and Denavit-Saubie, M., O2-sensing after carotid chemodenervation: Hypoxic ventilatory responsiveness and upregulation of tyrosine hydroxylase mRNA in brainstem catecholaminergic cells, Eur. J. Neurosci. 12, 3181–3190, 2000. 292. Coˆte´, A., Porras, H. and Meehan, B., Age-dependent vulnerability to carotid chemodenervation in piglets, J. Appl. Physiol. 80, 323–331, 1996. 293. Serra, A., Brozoski, D., Simeon, T., Yi, J., Bastasic, J., Franciosi, R., Jacobs, E.R. and Forster, H.V., Serotonin and serotonin receptor expression in the aorta of carotid intact and denervated newborns, Respir. Physiol. Neurobiol. 132, 253–264, 2002. 294. Sweeney, T.D., Leith, D.E. and Brain, J.D., Restraining hamsters alters their breathing pattern, J. Appl. Physiol. 70, 1271–1276, 1991. 295. Dauger, S., Nsegbe, E., Vardon, G., Gaultier, C. and Gallego, J., The effects of restraint on ventilatory responses to hypercapnia and hypoxia in adult mice, Respir. Physiol. 112, 215–225, 1998. 296. Kinkead, R., Dupenloup, L., Valois, N. and Gulemetova, R., Stress-induced attenuation of the hypercapnic ventilatory response in awake rats, J. Appl. Physiol. 90, 1729–1735, 2001. 297. Genest, S.E., Gulemetova, R., Laforest, S., Drolet, G. and Kinkead, R., Neonatal maternal separation alters development of the hypercapnic ventilatory response in awake adult rats (abstr), Respir. Res. 2(suppl 1), S31, 2001. 298. Kinkead, R. and Gulemetova, R., Neonatal maternal separation enhances time-dependent phrenic responses to hypoxia in rat (abstr), FASEB J. 17, A1297, 2003. 299. Schaefer, K.E., McCabe, N. and Withers, J., Stress response in chronic hypercapnia, Am. J. Physiol. 214, 543–548, 1968. 300. Ducsay, C.A., Fetal and maternal adaptations to chronic hypoxia: Prevention of premature labor in response to chronic stress, Comp. Biochem. Physiol. 119A, 675–681, 1998.
Respiratory Neuroplasticity
219
301. Challis, J.R., Sloboda, D., Matthews, S.G., Holloway, A., Alfaidy, N., Patel, F.A., Whittle, W., Fraser, M., Moss, T.J. and Newnham, J., The fetal placental hypothalamic-pituitary-adrenal (HPA) axis, parturition and post natal health, Mol. Cell. Endocrinol. 185, 135–144, 2001. 302. Joseph, V., Soliz, J., Pequignot, J., Sempore, B., Cottet-Emard, J.M., Dalmaz, Y., Favier, R., Spielvogel, H. and Pequignot, J.M., Gender differentiation of the chemoreflex during growth at high altitude: Functional and neurochemical studies, Am. J. Physiol. 278, R806–R816, 2000. 303. McEwen, B.S., Steroid hormones and the brain: Tinking ‘nature’ and ‘nurture,’ Neurochem. Res. 13, 663–669, 1988. 304. McEwen, B.S., How do sex and stress hormones affect nerve cells? Ann. NY Acad. Sci. 743, 1–16, 1994. 305. Simerly, R.B., Wired for reproduction: Organization and development of sexually dimorphic circuits in the mammalian forebrain, Annu. Rev. Neurosci. 25, 507–536, 2002. 306. Tatsumi, K., Hannhart, B. and Moore, L.G., Influence of sex steroids on ventilation and ventilatory control, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 829–864, 1995. 307. Schlenker, E.H. and Goldman, M., Acute effects of aspartic acid on ventilation in male and female rats, Physiol. Behav. 42, 313–318, 1988. 308. Shi, Y. and Schlenker, E.H., Neonatal sex steroids affect ventilatory responses to aspartic acid and NMDA receptor subunit 1 in rats, J. Appl. Physiol. 92, 2457–2466, 2002. 309. Schlenker, E.H., Goldman, M. and Holman, G., Effect of aspartic acid on control of ventilation in androgenized and ovariectomized female rats, J. Appl. Physiol. 72, 2255–2258, 1992. 310. Schlenker, E.H., Goldman, M. and Walsh, S., Perinatal estradiol benzoate administration affects control of ventilation in adult male rats, Physiol. Behav. 52, 1113–1116, 1992. 311. Ward, I.L. and Weisz, J., Differential effects of maternal stress on circulating levels of corticosterone, progesterone, and testosterone in male and female rat fetuses and their mothers, Endocrinology 114, 1635–1644, 1984. 312. Ramer, M.S., Harper, G.P. and Bradbury, E.J., Progress in spinal cord research: A refined strategy for the International Spinal Research Trust, Spinal Cord 38, 449–472, 2000. 313. Soden, R.J., Walsh, J., Middleton, J.W., Craven, M.L., Rutkowski, S.B. and Yeo, J.D., Causes of death after spinal cord injury, Spinal Cord 38, 604–610, 2000. 314. Claxton, A.R., Wong, D.T., Chung, F. and Fehlings, M.G., Predictors of hospital mortality and mechanical ventilation in patients with cervical spinal cord injury, Can. J. Anaesth. 45, 144–149, 1998. 315. Mansel, J.K. and Norman, J.R., Respiratory complications and management of spinal cord injuries, Chest 97, 1446–1452, 1990. 316. De Troyer, A., Estenne, M. and Vincken, W., Rib cage motion and muscle use in high tetraplegics, Am. Rev. Respir. Dis. 133, 1115–1119, 1986.
220
Fuller et al.
317. Haas, F., Axen, K., Pineda, H., Gandino, D. and Haas, A., Temporal pulmonary function changes in cervical cord injury, Arch. Phys. Med. Rehab. 66, 139–144, 1985. 318. Montero, J.C., Feldman, D.F. and Montero, D., Effects of glossopharyngeal breathing on respiratory function after cervical cord transection, Arch. Phys. Med. Rehab. 48, 650–653, 1967. 319. McDonald, J.W., Becker, D., Sadowsky, C.L., Jane, J.A., Sr., Conturo, T.E. and Schultz, L.M., Late recovery following spinal cord injury. Case report and review of the literature, J. Neurosurg. 97, 252–265, 2002. 320. Golder, F.J., Altered Respiratory Motor Output Following Cervical Spinal Cord Injury in Adult Rats, Ph.D. Dissertation, University of Florida, Gainesville, FL, 2002. 321. Goshgarian, H.G., The role of cervical afferent nerve fiber inhibition of the crossed phrenic phenomenon, Exp. Neurol. 72, 211–225, 1981. 322. Golder, F.J., Fuller, D.D., Davenport, P., Johnson, R.D., Reier, P.J. and Bolser, D.C., Respiratory motor recovery after unilateral spinal cord injury: Eliminating crossed phrenic activity decreases tidal volume and increases contralateral respiratory motor output, J. Neurosci. 23, 2494–2501, 2003. 323. Golder, F.J., Reier, P.J., Davenport, P.W. and Bolser, D.C., Cervical spinal cord injury alters the pattern of breathing in anesthetized rats, J. Appl. Physiol. 91, 2451–2458, 2001. 324. Golder, F.J., Reier, P.J. and Bolser, D.C., Altered respiratory motor drive after spinal cord injury: Supraspinal and bilateral effects of a unilateral lesion, J. Neurosci. 21, 8680–8689, 2001. 325. Nantwi, K.D., El-Bohy, A., Schrimsher, G.W., Reier, P.J. and Goshgarian, H.G., Spontaneous functional recovery in a paralyzed hemidiaphragm following upper cervical spinal cord injury in adult rats, Neurorehab. Neural Repair 13, 225–234, 1999. 326. Moreno, D.E., Yu, X.J. and Goshgarian, H.G., Identification of the axon pathways which mediate functional recovery of a paralyzed hemidiaphragm following spinal cord hemisection in the adult rat, Exp. Neurol. 116, 219–228, 1992. 327. Lipski, J., Zhang, X., Kruszewska, B. and Kanjhan, R., Morphological study of long axonal projections of ventral medullary inspiratory neurons in the rat, Brain Res. 640, 171–184, 1994. 328. Dobbins, E.G. and Feldman, J.L., Brainstem network controlling descending drive to phrenic motoneurons in rat, J. Comp. Neurol. 347, 64–86, 1994. 329. Rosenbluth, A. and Ortiz, T., The crossed respiratory impulses to the phrenic, Am. J. Physiol. 117, 495–513, 1936. 330. Porter, W.T., The path of the respiratory impulse from the bulb to the phrenic nuclei, J. Physiol. 17, 455–485, 1895. 331. Lindsay, A.D., Greer, J.J. and Feldman, J.L., Phrenic motoneuron morphology in the neonatal rat, J. Comp. Neurol. 308, 169–179, 1991. 332. Prakash, Y.S., Mantilla, C.B., Zhan, W.Z., Smithson, K.G. and Sieck, G.C., Phrenic motoneuron morphology during rapid diaphragm muscle growth, J. Appl. Physiol. 89, 563–572, 2000.
Respiratory Neuroplasticity
221
333. Guth, L., Functional plasticity in the respiratory pathway of the mammalian spinal cord, Exp. Neurol. 51, 414–420, 1976. 334. Chatfield, P.O. and Mead, S., Role of the vagi in the crossed phrenic phenomenon, Am. J. Physiol. 54, 417–422, 1948. 335. Lewis, L.J. and Brookhart, J.M., Significance of the crossed phrenic phenomenon, J. Neurophysiol. 166, 241–254, 1951. 336. Rosenbaum, H. and Renshaw, B., Descending respiratory pathways in the cervical spinal cord, Am. J. Physiol. 157, 468–476, 1949. 337. Goshgarian, H.G. and Guth, L., Demonstration of functionally ineffective synapses in the guinea pig spinal cord, Exp. Neurol. 57, 613–621, 1977. 338. Goshgarian, H.G., Developmental plasticity in the respiratory pathway of the adult rat, Exp. Neurol. 66, 547–555, 1979. 339. Ling, L., Bach, K.B. and Mitchell, G.S., Serotonin reveals ineffective spinal pathways to contralateral phrenic motoneurons in spinally hemisected rats, Exp. Brain Res. 101, 35–43, 1994. 340. Fuller, D.D., Johnson, S.M., Johnson, R.A. and Mitchell, G.S., Chronic cervical spinal sensory denervation reveals ineffective spinal pathways to phrenic motoneurons in the rat, Neurosci. Lett. 323, 25–28, 2002. 341. Johnson, R.A., Okragly, A.J., Haak-Frendscho, M. and Mitchell, G.S., Cervical dorsal rhizotomy increases brain-derived neurotrophic factor and neurotrophin-3 expression in the ventral spinal cord, J. Neurosci. 20, RC77, 2000. 342. Ling, L., Bach, K.B. and Mitchell, G.S., Phrenic responses to contralateral spinal stimulation in rats: Effects of old age or chronic spinal hemisection, Neurosci. Lett. 188, 25–28, 1995. 342a. Yu, X.J. and Goshgarian, H.G., Aging enhances synaptic efficacy in a latent motor pathway following spinal cord hemisection in adult rats, Exp. Neurol. 121, 231–238, 1993. 343. Kipnis, J., Yoles, E., Schori, H., Hauben, E., Shaked, I. and Schwartz, M., Neuronal survival after CNS insult is determined by a genetically encoded autoimmune response, J. Neurosci. 21, 4564–4571, 2001. 344. Hashimoto, T. and Fukuda, N., Contribution of serotonin neurons to the functional recovery after spinal cord injury in rats, Brain Res. 539, 263–270, 1991. 345. Heckman, C.J., Alterations in synaptic input to motoneurons during partial spinal cord injury, Med. Sci. Sports Exerc. 26, 1480–1490, 1994. 346. Saruhashi, Y. and Young, W., Effect of mianserin on locomotory function after thoracic spinal cord hemisection in rats, Exp. Neurol. 129, 207–216, 1994. 347. Saruhashi, Y., Young, W. and Perkins, R., The recovery of 5-HT immunoreactivity in lumbosacral spinal cord and locomotor function after thoracic hemisection, Exp. Neurol. 139, 203–213, 1996. 348. Zhou, S.Y., Basura, G.J. and Goshgarian, H.G., Serotonin(2) receptors mediate respiratory recovery after cervical spinal cord hemisection in adult rats, J. Appl. Physiol. 91, 2665–2673, 2001. 349. Mitchell, G.S., Bach, K.B., Martin, P.A., Foley, K.T., Olson, E.B., Brownfield, M.S., Miletic, V., Behan, M., McGuirk, S. and Sloan, H.E.,
222
350.
351.
352.
353.
354.
355.
356.
357. 358.
359.
360. 361.
362.
363.
Fuller et al. Increased spinal monoamine concentrations after chronic thoracic dorsal rhizotomy in goats, J. Appl. Physiol. 89, 1266–1274, 2000. Hadley, S.D., Walker, P.D. and Goshgarian, H.G., Effects of the serotonin synthesis inhibitor p-CPA on the expression of the crossed phrenic phenomenon 4 h following C2 spinal cord hemisection, Exp. Neurol. 160, 479–488, 1999. Zhou, S.Y. and Goshgarian, H.G., Effects of serotonin on crossed phrenic nerve activity in cervical spinal cord hemisected rats, Exp. Neurol. 160, 446–453, 1999. Zhou, S.Y. and Goshgarian, H.G., 5-Hydroxytryptophan-induced respiratory recovery after cervical spinal cord hemisection in rats, J. Appl. Physiol. 89, 1528–1536, 2000. Tai, Q., Palazzolo, K.L. and Goshgarian, H.G., Synaptic plasticity of 5-hydroxytryptamine-immunoreactive terminals in the phrenic nucleus following spinal cord injury: A quantitative electron microscopic analysis, J. Comp. Neurol. 386, 613–624, 1997. Sperry, M.A. and Goshgarian, H.G., Ultrastructural changes in the rat phrenic nucleus developing within 2 h after cervical spinal cord hemisection, Exp. Neurol. 120, 233–244, 1993. Hadley, S.D., Walker, P.D. and Goshgarian, H.G., Effects of serotonin inhibition on neuronal and astrocyte plasticity in the phrenic nucleus 4 h following C2 spinal cord hemisection, Exp. Neurol. 160, 433–445, 1999. Mantilla, C.B., Zielinska, W., Zhan, W.Z. and Sieck, G.C., C2 spinal cord hemisection alters Trk B, NT-4/5 and BDNF mRNA expression in rat cervical spinal cord and diaphragm muscle (abstr), FASEB J. 16, A771, 2002. Young, W., Fear of hope, Science 277, 1907, 1997. Nantwi, K.D. and Goshgarian, H.G., Effects of chronic systemic theophylline injections on recovery of hemidiaphragmatic function after cervical spinal cord injury in adult rats, Brain Res. 789, 126–129, 1998. Nantwi, K.D. and Goshgarian, H.G., Theophylline-induced recovery in a hemidiaphragm paralyzed by hemisection in rats: Contribution of adenosine receptors, Neuropharmacology 37, 113–121, 1998. Li, P. and Zhuo, M., Silent glutamatergic synapses and nociception in mammalian spinal cord, Nature 39, 695–698, 1998. Levine, E.S., Crozier, R.A., Black, I.B. and Plummer, M.R., Brain-derived neurotrophic factor modulates hippocampal synaptic transmission by increasing N-methyl-D-aspartic acid receptor activity, Proc. Natl. Acad. Sci. 95, 10235–10239, 1998. Baker-Herman, T.L., Fuller, D.D., Zabka, A.G., Johnson, R.A., Bavis, R.W. and Mitchell, G.S., Increased BDNF in the ventral cervical spinal cord following intermittent hypoxia requires spinal serotonin receptor activation and protein synthesis (abstr), Soc. Neurosci. Abstr. 27, 573.6, 2001. Doperalski, N.J., Fuller, D.D. and Mitchell, G.S., Chronic intermittent hypoxia (CIH), spinal hemisection and sham surgery alter cervical spinal brain derived neurotrophic factor (BDNF) concentration in rats (abstr), FASEB J. 17, A952, 2003.
Respiratory Neuroplasticity
223
364. Clark, G.A., Kandel, E.R., Induction of long-term facilitation in Aplysia sensory neurons by local application of serotonin to remote synapses, Proc. Natl. Acad. Sci. 90, 11411–11415, 1993. 365. Mitoma, H. and Konishi, S., Monoaminergic long-term facilitation of GABAmediated inhibitory transmission at cerebellar synapses, Neuroscience 88, 871–883, 1999. 366. Muir, G.D., Katz, S.L., Gosline, J.M. and Steeves, J.D., Asymmetric bipedal locomotion—An adaptive response to incomplete spinal injury in the chick, Exp. Brain Res. 122, 275–282, 1998. 367. Webb, A.A. and Muir, G.D., Compensatory locomotor adjustments of rats with cervical or thoracic spinal cord hemisections, J. Neurotrauma 19, 239–256, 2002. 368. El-Bohy, A.A., Schrimsher, G.W., Reier, P.J. and Goshgarian, H.G., Quantitative assessment of respiratory function following contusion injury of the cervical spinal cord, Exp. Neurol. 150, 143–152, 1998. 369. Wrathall, J.R., Pettegrew, R.K. and Harvey, F., Spinal cord contusion in the rat: Production of graded, reproducible, injury groups, Exp. Neurol. 88, 108–122, 1985. 370. Teng, Y.D., Mocchetti, I., Taveira-DaSilva, A.M., Gillis, R.A. and Wrathall, J.R., Basic fibroblast growth factor increases long-term survival of spinal motor neurons and improves respiratory function after experimental spinal cord injury, J. Neurosci. 19, 7037–7047, 1999. 371. Reier, P.J., Golder, F.J., Bolser, D.C., Hubscher, C., Johnson, J., Schrimsher, G.W. and Velardo, M.J., Gray matter repair in the cervical spinal cord, Prog. Brain Res. 137, 49–70, 2002. 372. Jammes, Y. and Speck, D.F., Respiratory control by diaphragmatic and respiratory muscle afferents, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 543–582, 1995. 373. Marlier, L., Poulat, P., Rajaofetra, N. and Privat, A., Modification of serotonergic immunoreactive pattern in the dorsal horn of the rat spinal cord following dorsal root rhizotomy, Neurosci. Lett. 128, 9–12, 1991. 374. Zhang, B., Goldberger, M.E. and Murray, M., Proliferation of SP- and 5HTcontaining terminals in lamina II of rat spinal cord following dorsal rhizotomy: Quantitative EM-immunocytochemical studies, Exp. Neurol. 123, 51–63, 1993. 375. Bach, K.B., Johnson, R.A., Kinkead, R., Fuller, D.D., Zhan, W., Mantilla, C., Sieck, G.C. and Mitchell, G.S., Cervical dorsal rhizotomy enhances serotonin-dependent long-term facilitation of hypoglossal motor output in rats (abstr), FASEB J. 14, A77, 2000. 376. Furicchia, J.V. and Goshgarian, H.G., Dendritic organization of phrenic motoneurons in the adult rat, Exp. Neurol. 96, 621–634, 1987. 377. Goshgarian, H.G., Ellenberger, H.H. and Feldman, J.L., Decussation of bulbospinal respiratory axons at the level of the phrenic nuclei in adult rats: A possible substrate for the crossed phrenic phenomenon, Exp. Neurol. 111, 135–139, 1991.
7 Airway Reflexes in Humans
TAKASHI NISHINO Chiba University Chiba, Japan
I.
Introduction
In general, a reflex consists of a neural receptor, afferent pathway, central synapses, efferent motor pathway, and effector organ; this general rule can be applied to airway reflexes (Figure 7.1). The effects of airway reflexes are very diverse and include changes in breathing pattern, maintenance of airway patency, defensive or protective reactions, alterations in bronchomotor tone, mucus-secreting response, and cardiovascular changes. These reflex responses are influenced by the nature and strength of the stimulant. In addition, some of these reflex responses are highly specific for the particular respiratory site. This chapter will deal chiefly with airway reflexes in humans, describing general characteristics and the function of airway reflexes, and some clinical problems related to anesthesia and respiratory care.
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Central nervous system (airway reflex program)
Afferent pathway
Afferent pathway
Nervous receptors
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Airway reflexes
Figure 7.1 Simplified model of airway reflex system.
II.
Central Nervous System
It is obvious that the central nervous system (CNS) plays a crucial role in initiation of airway reflexes. However, studies on the neurophysiology of airway reflexes have mainly defined the afferent pathways, leaving the CNS as a black box, and little is known about the central neural organization of airway reflexes. Since primary afferents from the receptors in the airways travel in the vagus (X), glossopharyngeal (IX), and trigeminal nerves (V), and converge in the solitary tract destined for synaptic contact with secondorder neurons in the nucleus tractus solitarius (NTS), the NTS must be viewed as the single most important site of early integration of afferent input relevant to the control of airway reflex systems. The NTS also is richly endowed with neuropeptides and other neuroactive substances [1–3] and recent studies have demonstrated the presence of various receptors in the central pathways such as opioid, serotonin, dopamine and N-methyl-D-aspartate (NMDA) receptors [4–6]. Although the study of CNS pharmacology modulating airway reflexes provides some important information, the brainstem neuronal network mechanisms that mediate airway reflexes are still poorly understood.
III.
Afferent Innervation of the Upper Airway and Receptors
The upper airway is composed of the nose, the pharynx, the larynx, and the extrathoracic portion of the trachea. The anatomy of the upper airway is very complex and its structural complexity reflects diverse functions such as
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phonation, olfaction, air conditioning, digestion, preservation of airway patency, and protection of the airways. Sensory nerve endings in and under the epithelium of the upper airway that respond to various stimuli are called upper airway receptors. Upper airway receptors can be classified into nasal, pharyngeal, laryngeal, and tracheal receptors, based on their location. We know little about the structure of these receptors although numerous afferent terminals have been described in the upper airways: free nerve endings, distributed among the epithelial cells, having either myelinated or non-myelinated fibers, as well as more organized structures like corpuscles and taste buds [7,8]. There are free nerve endings with myelinated afferent nerve fibers under the epithelium of the upper airways, which not only respond to a wide variety of irritant gases and aerosols but also show a rapidly adapting response to maintained mechanical deformation [9]. These nerve endings are especially sensitive to chemical and mechanical stimulation and are classified as irritant receptors [10] (Figure 7.2). With single-unit action potential recordings, Sant’Ambrogio et al. [11,12] showed in the larynx of the dog that in addition to irritant receptors, there are three groups of receptors classified on the basis of their responses to airflow and mechanical changes (Figure 7.3). These are: (1) cold (flow) receptors which are affected by changes in laryngeal temperature; (2) pressure receptors which are sensitive to changes in laryngeal transmural pressure, and (3) drive receptors which are affected by laryngeal motion. Although the extrapolation of such classification to other sites of the upper airways may not be entirely valid,
Cigarette smoke AP % CO2
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Figure 7.2 Responses of a laryngeal irritant receptor to cigarette smoke and distilled water. A.P.: action potentials. Exposure of the upper airway to smoke is indicated by the CO2 signal (Data from Ref. 10).
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Figure 7.3 Different types of laryngeal receptors responding to airflow and mechanical changes. (a) Behavior of laryngeal cold (flow) receptor during inhalation of warm air. Dog is spontaneously breathing through upper airway. Note that inspiratory modulation of receptor disappears when laryngeal temperature during inspiration is raised to expiratory level by inhalation of warm air (between arrows). A.P.: action potentials; Pes: esophageal pressure; V; airflow; Lar. Temp: laryngeal temperature. (b) Laryngeal pressure receptor responding to negative transmural pressure. Note that maximum activity is seen during upper airway occlusion, in which larynx is subjected to increased negative pressure. Recording from left SLN. A: upper airway breathing changed to tracheostomy breathing at arrow. B: upper airway breathing and upper airway occlusion. C: tracheostomy breathing and tracheal occlusion. (c) Laryngeal drive receptor responding to distortion caused by action of upper airway muscles. Abbreviations are the same as Figure 7.3(b). Note that an inspiratory activity is present and is not influenced by flow. Inspiratory activity increases when inspiratory drive increases (occluded efforts) (Data from Refs. 11 and 12).
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receptors in different sites of the upper airways have many properties in common. For example, an activity related to cooling of the nasal cavity has been demonstrated in the ethmoidal nerve of cats [13,14], rats [15], and guinea pigs [16]. Also, the presence of pressure-responsive receptors within the nasal passages has been demonstrated in the ethmoidal nerve of rats [17] and cats [14]. Although non-myelinated afferent fibers from the larynx have been described in the larynx of cats [18,19] and guinea pigs [20], they are thought to be scanty and information about their properties is apparently lacking. However, it has been reported that intralaryngeal capsaicin, given to stimulate C-fiber receptors, causes no respiratory response in dogs, but induces apnea and hypertension in rats [21]. Therefore, in some species, the functional role of C-fiber receptors may not be ignored.
IV.
Reflex Responses from the Upper Airway
Reflex activity in the upper airway is associated with two important respiratory functions subserved by the upper airway: (1) defense and protection of airway and (2) maintenance of airway patency.
A. Defense and Protection of Airway Nose
Mechanical and chemical irritation of the nasal mucosa can elicit various reflex responses. Although the sneeze is one of the representative respiratory reflexes from the nose, it has been studied far less than other nasal reflex responses. This is probably because the sneeze is easily blocked by anesthesia. Nevertheless, there is evidence to show that, in anesthetized cats, electrical stimulation of the anterior ethmoidal nerve, the posterior nasal nerve (PNN) or the infraorbital nerve (ION), all of which are branches of the trigeminal nerve, can elicit a sneeze identical to that induced by mechanical stimulation [22]. The trigeminal nerves carry most of the afferent fibers, which respond to chemical and mechanical irritation on the nasal mucosa. Therefore, the trigeminal nerve is considered the main afferent pathway for the sneezing reflex. However, strangely enough, the sneeze is inhibited when the ION is stimulated together with either the anterior ethmoidal nerve or the PNN [22], suggesting the presence of inhibitory pathways in the ION. Sneezing can also be elicited by local application of capsaicin, nicotine or formalin to the nasal mucosa, but higher doses of capsaicin that deplete substance P-containing nerves of the neuropeptide can prevent the sneeze caused by inhaled irritants [23].
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During sneezing, the respiratory changes are quite similar to those of coughing. Thus, the expulsive effort is preceded by a deep inspiration. Unlike the cough, however, the pharynx seems to be constricted and the forced expiration is via both the nose and the mouth. Bronchoconstriction is characteristically present with coughing, but occurs only infrequently with sneezing. While it is possible to perform voluntary coughing, it is virtually impossible to reproduce sneezing voluntarily. Stimulation of the nasal mucosa can also produce depression of breathing or even apnea (Figure 7.4). However, it is not known why the same stimulus may cause different reflex responses and what determines which reflex will occur. Depression of breathing and apnea are more commonly observed than sneezing, at least in experimental conditions, presumably because these reflexes are more resistant to general anesthesia than the sneeze reflex [24]. The apneic reflex, which is possibly related in mechanism to the diving reflex of aquatic animals, would prevent or limit penetration of water or irritant gases in the respiratory tract. Despite the fact that the nose is an important reflexogenic site in humans, depression
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Figure 7.4 Respiratory response to nasal insufflation of a pungent anesthetic (isoflurane) in a lightly anesthetized human. HR: heart rate; BP: arterial blood pressure; Ptr: intratracheal pressure; V_ : airflow; VT: tidal volume; PETCO2 : end-tidal PCO2: Fiso: isoflurane concentration.
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of breathing or apnea may be relatively insignificant in clinical situations. For instance, mechanical stimulation in the nose in the newborn rabbit can cause death [25], but this never happens in human babies. Larynx
The larynx is one of the most important reflexogenic sites. Defensive and protective reflexes originating from the larynx include cough, apnea, the expiration reflex, laryngeal closure, swallowing, bronchoconstriction, and mucous secretion. The richness of the reflex response is properly represented by the abundance of sensory endings in various laryngeal structures. Among various reflex responses, cough, the expiration reflex, and swallowing can be identified as defense reflexes that aim at removal of the irritant agent, while apnea, laryngeal closure and bronchoconstriction are described as protective reactions that aim at stopping harmful penetration of the noxious agent from the outside environment [26]. The reflex responses elicited from the larynx vary with the nature and strength of the stimulant. For example, weak mechanical and chemical stimulations may introduce only the laryngeal closure reflex [27,28]. With stronger stimuli, forceful expiratory efforts such as the expiration reflex and coughing may be induced. The expiration reflex consists of a brief expiratory effort not preceded by an inspiration [29]. In contrast, the cough reflex usually starts with a brief rapid inspiration, followed immediately by a compressive phase in which the glottis is closed and the expiratory muscles contract forcefully together with the diaphragm [30]. An expulsive phase follows immediately, when the glottis is suddenly reopened, allowing the expulsion of a blast of air. During that time, oscillation of tissue and gas causes a characteristic explosive sound and may play a role in suspending secretions in the moving gas stream. Coughing elicited from the larynx appears to be similar in its fundamental characteristics to that evoked from the trachea. However, it has been shown in animal studies that the pattern of coughing from the larynx is slightly different from that induced from the trachea (laryngeal coughing has a higher frequency, a paroxysmal nature and stronger inspiratory efforts) [30]. Although there is no doubt that the larynx is an important tussigenic area, several studies raised the question about the importance of laryngeal afferent input in the cough reflex. Stockwell et al. [31] showed that anesthetic block of the superior laryngeal nerve (SLN) in man did not affect the cough threshold to inhaled nebulized citric acid in awake humans, suggesting that SLN afferents do not play a necessary role in initiation of citric acid-induced cough. A similar conclusion may be derived from the study of Higenbottam et al. [32] who showed that patients with cardiopulmonary transplants showed a poor or absence cough response to distilled water aerosols. It seems that coughing due to inhaled irritants is not
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much changed by laryngeal denervation; however, this does not necessarily mean that the larynx cannot be an important site for the initiation of cough. Pharynx
Mechanical stimulation of the mucosa of the nasopharynx in cats can elicit a sniff-like aspiration reflex that produces repeated powerful contraction of the diaphragm [33]. The purpose of this reflex response may be removal of foreign materials from the back of the nose into the pharynx to be swallowed or expelled by coughing. Korpas and Tomori [30] have shown that this reflex is exceptionally resistant to anesthesia, hypercapnia, asphyxia, hypothermia, and antitussive drugs. Although this reflex has been demonstrated in many other mammalian species, it seems that it is very weak in humans, including in neonates. The most obvious reflex response elicited from the pharynx is reflex swallowing. Although the main function of reflex swallowing is the propulsion of food from the oral cavity into the stomach, it also can serve as a protective reflex for the respiratory tract [34]. In general, swallowing can be divided into three states: (1) the initial oral preparatory state, (2) the subsequent pharyngeal state, and (3) the esophageal state. Of the three states, the involuntary control of the pharyngeal state of swallowing is the most important state from the standpoint of airway protection. Swallowing results in reflex closure of the glottis, the single most vital function of the larynx. Strong adduction of the true vocal cords is supplemented by the closure of the false cords and approximation of the aryepiglottic folds, although adduction of the true cords alone suffices to prevent entrance into the trachea of swallowed material. Swallowing must interact with respiration so that a swallow causes minimal or no disturbance of continual respiration [35]. In awake human adults, approximately 80% of swallows occur during the expiratory phase, and respiratory movement resumes after a swallow in the same expiratory phase as has been interrupted [36]. The preponderant coupling of swallows with the expiratory phase may be a useful mechanism for clearing the airway of foreign materials before the subsequent inspiration and thus may exert a physiologic role in preventing low-grade recurrent aspiration. This coupling of swallows with the expiratory phase is lost in unconscious adults, suggesting that the mechanism responsible for the preferred timing of swallows during the expiratory phase may have both centrally programmed and learned behavioral components [37]. The preponderant coupling of swallows with the expiratory phase is also lost during hypercapnia [38] and with the addition of respiratory elastic loading in conscious humans [39]. In these conditions, the timing of swallows shifts from the expiratory phase towards the inspiratory phases of the next breath. It may be possible that with this shift, incompletely eliminated pharyngeal content is aspirated into the
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larynx during the subsequent inspiration. In fact, it has been suggested that swallows coinciding with the expiratory–inspiratory transition phase are the most liable to produce aspiration [40].
B. Maintenance of Airway Patency
Until recently, only the defensive role of upper airway reflexes had been studied in detail, although it has been known for many years that pressure and airflow in the upper respiratory tract can change breathing [41]. The results of several studies [42–48] that analyzed the mechanisms of sleep apnea and sudden infant death syndrome indicate that upper airway pressure reflexes have important actions on the musculature of the pharynx and therefore on its patency, on breathing, and on arousal. For example, negative pressure in the upper airway produces an excitatory effect on upper airway dilating muscles while exerting an inhibitory effect on breathing patterns by prolonging inspiratory and expiratory durations and by decreasing the rate of rise of diaphragmatic activity [42–44,47,48]. The main reflexogenic site responding to pressure changes is thought to be the larynx that is supplied by the internal branch of the SLN, whereas the nose plays a subsidiary role and the oropharynx does not seem to be important. Thus, the negative upper airway reflex can be abolished by mucosal application of topical anesthetics or by cutting the SLN [43,49]. Both the excitatory influences on the upper airway controlling muscles and the inhibitory influences on chest wall muscles can be interpreted as contributing to airway stability in that they increase the dilating forces and reduce the collapsing forces, respectively. In fact, the elimination of sensory feedback from the upper airway impairs the ability of upper airway muscles to respond to adverse conditions such as airway obstruction [50]. Other studies [51,52] also showed that in awake humans, a strong activation of the genioglossus muscle occurs in response to negative pressure applied in the upper airway, which can be abolished by upper airway anesthesia. Although application of negative pressure causes genioglossus muscle activation in both wakefulness and sleep, the increase in genioglossus activity is attenuated during non-Rapid Eye Movement (non-REM) sleep in adult subjects as well as in neonates [53,54]. Regarding maintenance of airway patency, the ability to switch the breathing route appears to be another important function. Acute obstruction of nasal passages leads to opening the mouth and oral breathing. The ability to change from the nasal route of airflow to the oral route during nasal obstruction is crucial for maintenance of adequate ventilation. Two major muscles responsible for the switching of breathing route are the palatoglossus muscle, which directs the soft palate caudally and ventrally, and the levator veli palatini, which pulls the soft palate cephalad and in
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a dorsal direction. Activation of these muscles results in nasal breathing and oral breathing, respectively. Consciousness is an important factor in the control of the route of breathing [55], but the switching of breathing route could also be triggered by some reflex mechanisms. It has been shown that in awake subjects, application of topical lidocaine in the nasal passages delays the onset of oral breathing in response to nasal occlusion (Figure 7.5), suggesting that in humans, sensory information from receptors in the nasal passage has an important role in controlling the shift of breathing route [56]. V.
Afferent Innervation of the Lower Airway and Receptors
The vagus nerve and its bronchial branches innervate the tracheobronchial tree and lung parenchyma. In contrast to the dense parasympathetic nerve supply to the lower airways, adrenergic innervation is sparse in humans [57]. Furthermore, there is no evidence for functional sympathetic innervation in isolated human smooth muscle [58]. It is generally agreed that in the lower airway, there are at least four different types of receptors: (1) slowly adapting receptors (SARs); (2) rapidly adapting receptors (RARs); (3) C-fiber endings, and (4) sensory receptors in neuroepithelial bodies (NEBs). Of these, the structure of only two types of receptor, i.e., SARs and NEBs, can be stated with confidence despite numerous studies on the morphology of the sensory innervation of the airways. The reflex function of the latter is unclear [59].
Nasal chamber pressure (cm H2O) Mouth chamber pressure (cm H2O) SaO2 (%)
5 0 1 0
BEFORE ANAESTHESIA
AFTER ANAESTHESIA
100 80
End-tidal CO2 nose (mm Hg)
40 0
End-tidal CO2 mouth (mm Hg)
40 0
10 S
Figure 7.5 Respiratory responses to nasal occlusion in one nasal breather. The subject was breathing through a face mask, which separates nasal and oral passages. Horizontal bars indicate nasal obstruction and arrows indicate the beginning and end of oral breathing. Note that oral breathing starts immediately after nasal obstruction before topical nasal anesthesia whereas the start of oral breathing delays considerably after topical nasal anesthesia (Data from Ref. 56).
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A. Slowly Adapting Receptors (SARs)
Slowly adapting receptors are found in the smooth muscle of the larger airways and correspond to the myelinated afferent nerve fibers in the vagus. The SARs are the more easily identifiable of the endings, having a respiratory modulation because of the regular characteristics of their discharge both during transient changes in lung volume and during maintained inflations. Some SARs have a continuous discharge during the respiratory cycle with an increasing rate during inspiration, indicating the activation threshold of these receptors is below functional residual capacity (FRC), whereas others can fire only above FRC [60] (Figure 7.6). In general, they are not chemosensitive, but they are stimulated by drugs such as histamine and acetylcholine, which contract smooth muscle [61,62]. Inhalation of CO2 and volatile anesthetics is also known to affect the
(a)
(b)
A.P.
Ptr
cmH20 5 0 1s
(c)
(d)
A.P.
cmH20 Ptr
5 0
Figure 7.6 Subtypes of pulmonary stretch receptors (SARs). (a) Low-threshold SARs with tonic expiratory discharge; (b) low-threshold SAR with cardiac modulation; both (c) and (d) are high-threshold SARs, but tracheal threshold pressure of (c) is much higher than that of (d) (Data from Ref. 60).
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activity of SARs [63]. Inhalation of CO2 inhibits their activity by a direct action on the receptor with action on 4-aminopyridine-sensitive Kþ channels [64], whereas volatile anesthetics may inhibit or stimulate the receptors, depending on the concentration and the type of SAR [60]. B. Rapidly Adapting Receptors (RARs)
Although the structure of RARs has not been fully delineated, RARs are known to have non-myelinated terminals connected to thin myelinated vagal afferents (A) [59,63]. They are found throughout the lower airways but are scanty in the smaller bronchi, and none has been identified in the bronchioles and alveoli. As the name implies, these receptors adapt rapidly to a maintained inflation or deflation of the lungs. The respiratory modulation of the RARs is irregular in both its timing with the breathing cycle and its pattern of discharge. RARs are activated by a large number of mechanical and chemical irritant stimuli (ammonia, ether vapor, cigarette smoke, etc.), by inflammatory and immunological mediators, and by airway and lung pathological changes. All the stimuli that can induce coughing can also stimulate RARs, and therefore, it is likely that RARs are directly involved in elicitation of the cough reflex [65]. Also, most of the mechanical and chemical irritants that stimulate RARs are effective bronchoconstrictors, although cough and bronchoconstriction are two separate reflexes [32]. C. C-fiber Endings
Two groups of C-fiber receptors have been distinguished on the basis of their circulatory accessibility through either the pulmonary or the bronchial circulation [66,67]. Pulmonary C-fiber receptors are those arising from the endings located in the lung parenchyma, and are directly accessible to a challenging drug injected into the pulmonary artery, whereas bronchial C-fiber receptors located further down stream, innervating the airway mucosa, are accessible to the challenging drug injected into the left atrium or directly into the bronchial artery (Figure 7.7). Although the distinction between pulmonary and bronchial C-fiber receptors was based initially on the location and circulatory accessibility, the two have been found to differ somewhat in their afferent properties. For example, pulmonary C-fiber receptors show a relatively greater mechanosensitivity than do bronchial C-fiber receptors in dogs [67,68]. Pulmonary C-fiber endings are relatively insensitive to autacoids such as bradykinin, histamine, serotonin, and prostaglandins, whereas bronchial C-fiber endings are sensitive to a wide range of intrinsic chemicals including histamine, bradykinin, and prostaglandins, either injected into the bronchial artery or administered as aerosol
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Pulmonary C-fiber and rapidly adapting receptor
5 sec.
ENG Pes 0 (cmH20) −10 0 Ptr −10 (cmH20) −150 100 BP (mmHg) 50
PTO2
21 16 5 0
(%)
PTCO2 (%)
Marker
Inj.
Bronchial C-fiber ENG Pes
0
(cmH20) −10 Ptr 0 (cmH20) −10 −150
BP 100 (mmHg) 50 PTO2 (%)
PTCO2 (%)
21 16 5 0
Marker Inj.
Figure 7.7 Responses of pulmonary C-fiber (small spikes) and rapidly adapting receptor (larger spikes) (upper panel) and a bronchial C-fiber (lower panel) to the right atrial injection of capsaicin (10 mg/kg) in an anesthetized, spontaneously breathing dog. Note that the pulmonary C-fiber was activated immediately after the capsaicin injection, but the bronchial C-fiber had a longer latency, whereas in both cases apnea was followed by hypotension and bradycardia. The pulmonary C-fiber was located in the central part of the left lower lobe and the bronchial C-fiber ending was located at the hilum of the left lower lobe. ENG: electroneurogram; Pes: esophageal pressure; Ptr: intratracheal pressure, BP: arterial blood pressure; PTO2 : PO2 in tidal air; PTCO2 : PCO2 in tidal air. The injection time is marked on the bottom trace (Data from Ref. 66).
[67–69]. In contrast, the two groups of C-fiber receptors respond similarly to inhalation of volatile anesthetics [66]. D. Neuroepithelial Bodies (NEBs)
The airway and alveolar epithelia contain pulmonary neuroendocrine cells that contain a large range of neuroendocrine markers and bioactive substances such as serotonin, calcitonin gene-related peptide, and the mitogen bombesin. These cells are sometimes collected into clusters called neuroepithelial bodies (NEBs). The NEBs are innervated and the recent evidence shows that there are two separate populations of sensory nerve fibers that selectively contact the NEBs, i.e., the vagal sensory component
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and the spinal sensory component [70]. The NEBs are thought to be sensors of hypoxia, and hypoxia-sensitive Kþ channel have been identified in their membranes [71]. Hypoxia is presumed to release mediators from the NEBs, which in turn stimulate the sensory nerves and exert local actions on airway smooth muscle and the bronchial and pulmonary vascular beds [59]. However, there is no evidence to suggest that sensory nerves associated with NEBs are involved in elicitation of airway defensive reflexes. VI.
Integrative Aspects of the Airway Reflexes Elicited from the Lower Airways
A. Lung Inflation and Activation of SARs
It is generally believed that inflation of the lungs activates SARs and causes the classical Breuer-Hering inflation reflex that provides the predominant mechanism for regulating the depth and rate of breathing in anesthetized animals. Although there are some reports to suggest that the Breuer-Hering inflation reflex is operative at normal tidal volume in human subjects [72,73], this reflex is considered very weak in healthy eupneic man, particularly in conscious conditions. Hamilton et al. [74] studied the effect of passive lung inflation during wakefulness and during stable non-REM sleep in laryngectomized patients. During sleep, apnea, as the evidence of the Breuer-Hering inflation reflex, was produced by lung inflations only when inflation volumes exceeding 1 liter were applied at end-inspiration, whereas no change in respiratory timing was apparent during wakefulness even with greater inflation volume, suggesting that the Breuer-Hering inflation reflex can be demonstrated above the resting tidal volume range in adult man only in the absence of the behavioral control of breathing. In contrast to the responses observed during lung inflation, deflation of the lungs causes reflex tachypnea and a reflex increase in inspiratory drive in most species including man [75]. Considering the experimental results that deflation reduces the input from SARs and stimulates RARs [76,77], the reflex excitatory effect on breathing appears to result from stimulation of RARs. Inflation of lungs causes not only Breuer-Hering inflation reflex but also reflex relaxation of smooth muscle [78]. This reflex is predominantly attributable to inhibition of parasympathetic cholinergic motor tone since atropine can prevent this reflex action. Although clinical significance of this reflex is not entirely clear, it has been proposed this reflex could adjust airway geometry in different patterns of breathing to optimize the relationship between airways resistance and deadspace [79]. The significant role of SARs on coughing was implicated first by Bucher [80] who suggested that during the inspiratory effort, which is an
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integral part of the cough reflex, SARs are more intensively stimulated, thereby increasing the inhibitory influence on central inspiratory activity and thus strengthening the subsequent expiration. The studies of Hanacek et al. [81] and Sant’Ambrogio et al. [82] showed that in anesthetized rabbits, stimulation of tracheal mucosa failed to elicit cough when high concentrations of SO2 had abolished the Breuer-Hering inflation reflex (Figure 7.8). In anesthetized humans, lung inflation with CPAP of 10 cmH2O did not exert any influence on the reflex responses to tracheal irritation in terms of the types, latencies, and duration of reflex responses, suggesting the role of SARs in elicitation of cough is speculative in anesthetized humans [83]. However, information obtained in anesthetized subjects may not be entirely applicable to awake subjects. Since it has been reported that the cough response to inhalation of nebulized distilled water is remarkably diminished in awake, heart-lung transplantation patients whose lungs are denervated below the level of the tracheal anastomosis, compared with normal subjects [32], it is possible that loss of the facilitatory influence of SARs on
Control BP
SO2
100
(mmHg) 0 Dia. EMG Abd. EMG 10 Pes 0 (cmH2O) –10 PLC 10 0 (cmH2O)
BP
100
(mmHg) 0 Dia. EMG Abd. EMG Pes 10 (cmH2O) 0 –10
Figure 7.8 Effects of SO2 administration on the cough reflex in an anesthetized rabbit. In the control situation, lung inflation to 10 cmH2O inhibited breathing (Hering-Breuer inflation reflex). Cough (indicated by increased activity in diaphragm and abdominal muscles) could be elicited by tracheal stimulation. After SO2 administration (200 ppm), the Hering-Breuer inflation reflex has disappeared and cough could not be elicited with tracheal stimulation. Left-hand tracings: before SO2 administration; right-hand tracings: after SO2 exposure. Upper panels: effects of lung inflation. Lower panels: effects of tracheal stimulation (Data from Ref. 82).
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coughing could contribute to the diminished cough response during laryngeal irritation in these patients. B. Reflex Responses Elicited by Mechanical and Chemical Stimulation of the Lower Airways
Considerable differences exist among species with respect to the type and magnitude of reflex responses elicited from the lower airways. For example, mice and ferrets show no cough response to mechanical stimulation of the tracheal mucosa [84,85] whereas the same stimulation elicits vigorous coughing in cats and dogs [86,87]. The reflex responses evoked from the stimulation of the lower airways are linked to arousal. Sullivan et al. [88] demonstrated that a stimulus sufficient to cause coughing during wakefulness failed to do so in either SWS or REM sleep in the absence of arousal. When the stimulus was sufficient to cause arousal, coughing always followed arousal, suggesting that coughing involves supra-medullary neural processes that are normally active only during wakefulness. In anesthetized animals, irritation of the trachea with mechanical stimuli causes coughing, hypertension and laryngeal constriction, whereas irritation of the bronchi with chemical stimuli causes hyperpnea [86,87]. Also, in general the trachea and its vicinity are very sensitive to mechanical stimulation whereas the bronchi are more sensitive to chemical stimulation. These differences in the reflex responses seem to depend on the difference in the characteristics of RARs in different sites, since it is known that RARs in the trachea and larger bronchi are very mechanosensitive whereas RARs in bronchi are more chemosensitive [59]. In anesthetized humans, at least six different types of respiratory responses are observed during stimulation of the tracheal carina induced by injection of a small amount of distilled water [89]: namely (1) the apneic reflex, (2) the expiration reflex, (3) the cough reflex, (4) spasmodic panting, (5) slowing of breathing, and (6) rapid, shallow breathing. However, the level of anesthesia has a marked influence on these reflex responses. Thus, among these reflex responses, the cough reflex is the most sensitive and the apneic reflex is the most resistant to deepening anesthesia, whereas the other types of reflex responses were in between (Figure 7.9). Airway receptors responsible for eliciting airway reflexes do not appear to be uniformly distributed in the airways [62], and therefore it is not surprising to observe the differences in the reflex responses from different sites in the airways. In contrast with the reflex responses elicited from the tracheal carina, the same stimulation given to the bronchus causes little or no reflex response [90] (Figure 7.10). This observation is in agreement with the observation made by Jackson [91] that a small mechanical irritant in the trachea causes vigorous coughing whereas mechanical stimulation of the
Airway Reflexes in Humans
% RESPONSES
100
241 Expiration reflex
Apnea
Spasmodic, panting breathing
80 60 40 20 1.3
% RESPONSES
100
1.0
0.7 MAC
Slowing of breathing
Cough reflex
80
Rapid, shallow breathing
60 40 20
1.3
1.0
0.7 MAC
Figure 7.9 Occurrence of various respiratory responses to injection of distilled water into the trachea at three different depths of enflurane anesthesia in humans. Ordinate, percent of positive responses. MAC: minimum alveolar concentration (Data from Ref. 89).
finer subdivisions of the tracheobronchial tree causes less cough production during bronchoscopic procedures in awake patients, suggesting the peripheral bronchial branches are less important as reflexogenic areas in humans. Cigarette smoke is one of the most common inhaled irritants in human airways and is known to evoke coughing and reflex bronchoconstriction. Although it has been reported that cigarette smoke activates RARs in the larynx and tracheobronchial trees and thereby elicits coughing and bronchoconstriction [26], the study of Lee et al. [92] demonstrated that in dogs, inhaled cigarette smoke triggers a consistent and vigorous stimulation of pulmonary C-fiber afferents. Nicotine is considered to be primarily responsible for triggering this response, since the response is completely abolished by pretreatment with hexamethonium, a nicotinic acetylcholinereceptor antagonist. In agreement with this idea, in healthy human nonsmokers, the intensity of cigarette smoke-induced airway irritation and cough responses following cigarette smoke inhalation are greatly attenuated by premedication with aerosolized hexamethonium [93] (Figure 7.11).
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(bpm) HR 120 80 (mmHg) BP 120 80 (cmH2O) AP 5 0 (1/min) AF 0.5 0
VT
l 0.3
0 (mmHg) 40 PET CO2 0
(bpm) HR 120 80 (mmHg) BP 120 80 (cmH2O) AP 5 0 (1/min) AF 0.5 0 VT
l 0.3
0 (mmHg) 40 PET CO2 0
Figure 7.10 Respiratory responses to stimulation by distilled water (given at arrows) of trachea and bronchus. Upper panel: tracheal stimulation; Lower panel: bronchial stimulation. Note that stimulation of trachea causes vigorous reflex responses of respiration and circulation whereas bronchial stimulation caused no reflex response. HR: heart rate; BP: arterial blood pressure; AP: airway pressure; AF: air flow; VT: tidal volume; PETCO2 : end-tidal PCO2 (Data from Ref. 90).
Ozone is another potentially harmful chemical stimulus with irritant effects on the lower airways. In dogs, inhalation of ozone into the lower trachea causes tachypnea and bronchoconstriction [94]. In human subjects, breathing ozone for even relatively short periods causes rapid shallow
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Airway irritation (total number of push-button signals)
After placebo 10
After hexamethonium 10
10
8
8
8
6
6
6
4
4
4
2
2
2 0
0
0 0-5
5-15
15-30
Time after smoke inhalation (sec)
Figure 7.11 Comparison of responses to inhalation of high-nicotine cigarette smoke after premedication with aerosolized saline placebo and hexamethonium (10%) (Data from Ref. 93).
breathing, cough, inspiratory chest pain, and bronchoconstriction [95,96]. These reflex responses are thought to be mediated by the increase in vagal C-fiber receptors in the lower airways. In fact, there is evidence to show that the sensitivity of bronchial C-fiber receptors can be increased by ozone in dogs [97] and rats [98]. However, ozone can also activate RARs [97]. Therefore, it is unlikely that all the ozone-induced reflex responses are mediated by bronchial C-fiber receptors. Although a predominant role of C-fiber receptors in the changes of breathing pattern, such as apnea and rapid shallow breathing in response to extraneous chemicals, is well recognized, the role of airway C-fibers in initiating cough remains uncertain and has been the subject of considerable debate. Inhalation of capsaicin aerosol reproducibly causes cough in humans [99] and conscious guinea pigs [100]. In addition, tachykinins, especially substance P, can induce coughing in guinea pigs [101]. In normal and asthmatic subjects, substance P aerosols cause no cough [102], but this agent induces cough in patients with upper respiratory tract infections [103]. By contrast, systemic administration of capsaicin does not usually elicit cough [26] with an exception in one study with conscious humans [104]. Furthermore, there is experimental evidence to show that C-fiber activation can inhibit cough in anesthetized animals [105] (Figure 7.12). In addition, capsaicin is not sufficiently specific as a C-fiber stimulant, since capsaicin can stimulate not only C-fiber endings but also the endings of RARs [106].
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(b)
EMG
0.4 V′ (l.s-1)
0.2 0 0.2 0.4
BP (kPa)
20 10 0
30 s
Figure 7.12 Effect of pulmonary C-fiber reflex on coughing induced from the tracheobronchial tree of unanesthetized cat. EMG: electromyographic activity of genioglossus muscle; V 0 : airflow from tracheal cannula; BP: arterial blood pressure. (a) The tracheobronchial mucosa was stimulated mechanically during the signal marks, causing increased EMG activity and airflow, corresponding to cough efforts. The cough efforts continued long after the stimulus had stopped. (b) Phenylbiguanide (25 g/kg) was injected intravenously at the arrow, causing hypotension, bradycardia and apnea due to stimulation of pulmonary C-fiber receptors. During the apnea, the tracheobronchial stimulus was repeated at the signal marks, causing no change in airflow but some increase in EMG activity. Later, during the phase of rapid shallow breathing, the tracheobronchial stimulus was repeated and caused cough efforts, with no coughing after the end of stimulus (Data from Ref. 105).
Thus, it has been argued that inhaled capsaicin could cause cough by stimulating RARs, either directly or indirectly by releasing tachykinins from C-fiber endings, which sensitizes the endings of RARs, or by initiating reflex bronchoconstriction that in turn stimulates the RARs [59,65] (Figure 7.13). VII.
Clinical Problems Associated with Airway Reflexes
A. Pulmonary Aspiration
The pulmonary aspiration of oropharyngeal or gastric contents into the lower respiratory tract is a major complication observed during perioperative periods and a life-threatening danger to every comatose and debilitated
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C-fiber receptor
RAR Capsaicin Irritants, etc.
Mucus Mechanical Epithelium
Tachykinins
Mucosa
Peptidases Tachykinin Antagonists Antipeptidases Inhibit cough
CNS
Excite cough
Cough
Figure 7.13 Diagram of hypothetical role for tachykinins in cough. Tachykinins may be released from C-fiber receptors, and diffuse to RARs, which they stimulate causing cough. They can be broken down by peptidases, which in turn can be inhibited by antipeptidases. Tachykinin antagonists can prevent the action of tachykinins on the RARs. If there is sufficient stimulation of C-fiber receptors, these can cause a central inhibition of cough. CNS: central nervous system; RAR: rapidly adapting receptor (Data from Ref. 65).
patient [34,107]. Several pulmonary syndromes including aspiration pneumonitis, aspiration pneumonia, airway obstruction, lung abscess and chronic interstitial fibrosis may occur after aspiration, depending on the amount and nature of the aspirated material, the frequency of aspiration, and the host’s response to the aspirated material [107]. Among these pulmonary syndromes, aspiration pneumonitis and aspiration pneumonia are the two main syndromes, and although there is some overlap between these syndromes, they are distinct clinical entities. Aspiration pneumonitis, known as acid aspiration syndrome or Mendelson’s syndrome, is a chemical lung injury that occurs in patients who have a marked disturbance of consciousness such as resulting from stroke, drug overdose, or general anesthesia. On the other hand, aspiration pneumonia is an infectious process caused by the inhalation of oropharyngeal secretions that are colonized by pathogenic bacteria. Reliable incidences of clinical aspiration pneumonitis and aspiration pneumonia are difficult to determine since the epidemiologic study of aspiration syndromes is not sufficient. It has been reported that aspiration
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pneumonitis occurs in approximately 10% of patients who are hospitalized after drug overdose [108]. A multicenter, prospective study of nearly 200,000 operations in France from 1978 to 1982 found the overall incidence of aspiration to be 1.4 per 10,000 anesthetics [109]. A review of computer-based records of approximately 185,000 anesthetics administered to adults from 1967 to 1983 in Sweden noted an incidence of 4.7 aspirations per 10,000 anesthetics [110]. The incidence of aspiration pneumonia is more difficult to determine. Several studies indicate that 5–15% of cases of communityacquired pneumonia are aspiration pneumonia [111–113]. There is no doubt that defensive airway reflexes, including the swallowing reflex, play a crucial role in prevention of pulmonary aspiration [34,107]. On the other hand, there are certain clinical situations where a suppression of defensive airway reflexes is desirable. For example, the SLN block may be the choice of technique to facilitate diagnostic laryngoscopy and bronchoscopy as well as to allow the comfortable placement of a tracheal tube in those patients in whom a difficult intubation is anticipated, since this technique can minimize the adverse effects of upper airway reflexes during airway maneuvers [114]. Impairment of defensive airway reflexes may result from a defect or disorder in any part of the reflex arc of the defensive airway reflexes shown in Figure 7.1. Since afferent nerve endings are the natural starting of all reflex activity, it is natural that impairment of triggering defensive airway reflexes occurs after application of local anesthetics into the upper airway. As mentioned above, nerve blocks of the afferent pathway with local anesthetics are useful in elimination of various defensive airway reflexes. The impairment of defensive airway reflexes may also occur in the CNS. It is a common observation that depression of defensive airway reflexes can occur not only during general anesthesia but also during sedation in surgical patients [115,116]. Dysfunctions of efferent neural pathways and effector organs may also seriously impair the defensive airway reflexes. Muscle disorders, disorders of the neuromuscular junction, and disorders that affect peripheral nerves all can lead to impairment of reflex responses and may result in pulmonary aspiration [117]. B. Cough
Coughing is very rare in complete health. On the other hand, chronic cough is one of the most common symptoms seen in ambulatory practice [118]. It is also well recognized that cough is associated with many kinds of diseases such as upper airway infections, asthma, bronchitis, and pneumonia as well as allergy, various occupational exposures, and cigarette smoking [119]. Thus cough is an index of disease and can be used as a tool in differential diagnosis. Although cough is a normal protective reflex, in disease it may be
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persistent, impairing bodily functions and becoming an embarrassment [120]. For example, distal bronchial secretions tend to be spread peripherally during cough and may spread disease within the lung. Trauma to the tracheobronchial wall or larynx may lead to bleeding or may play a role in the development of infection. The muscular effort involved in coughing is relatively great and may aggravate heart failure in patients who have passive congestion of the lungs. Ribs may be fractured as the result of cough [121], and this is a not uncommon cause of severe chest pain. Muscle soreness likewise develops in chronic prolonged cough. During a prolonged paroxysm of cough, the persistent high intra-thoracic pressure may so impede venous return that cardiac output falls and cerebral ischemia occurs [122]. This form of cough syncope may be associated with fainting and convulsions, and is most common in the elderly with cerebral arteriosclerosis. Chronic cough is associated with the risk of myocardial infarction [123]. Cough may also be harmful in a public health sense because of its potential for spreading infections. When the cough is troublesome and serves no useful purpose, the cough has to be treated effectively. The rational clinical approach to the management of chronic cough is to search for an underlying cause and to treat the cause. This approach is usually easy and successful in the majority of patients with chronic cough. Indeed, Irwin et al. [124] showed that utilizing the anatomic, diagnostic protocol, the causes of cough were determined in 99% of their patients, leading to specific therapy that was successful in 98%. It has been reported that the cough reflex in patients with chronic cough is sensitized in several associated conditions, including angiotensin-converting enzyme (ACE) inhibitor cough, gastroesophageal reflux (GER), and cough-variant asthma [125]. However, the mechanism responsible for sensitization of the cough reflex remains obscure. Angiotensin-converting enzyme inhibitor cough occurs in 0.2–33% of patients treated with ACE inhibitors [126], recurring with reintroduction of the same or another ACE inhibitor. It is more common in women. The mechanism may involve accumulation of endogenous tachykinins or bradykinin in the airway due to prevention of breakdown of kinins by ACE, which may stimulate RARs and induce coughing [65,127] (Figure 7.13). Gastroesophageal reflux is a common cause of chronic cough [124,128]. Although some patients may have reflux up to the pharynx and may even aspirate, it is unlikely that aspiration of acid reflux may be a prerequisite. There is evidence to indicate that the cough reflex can be caused by activation of sensory receptors in the distal esophagus [129]. Cough-variant asthma is an occult form of asthma of which the only sign or symptom is chronic cough [130,131]. The diagnosis of cough-variant asthma is made only by demonstration of airway hyperresponsiveness to methacholine or histamine challenge and an excellent response to treatment
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with bronchodilators and corticosteroids [132]. Why some patients with asthma have cough as the principal feature of their disease is totally unclear. Postnasal drip syndrome (PNDS), either singly or in combination with other conditions, is the single most common cause of chronic cough in nonsmoking patients [124]. Although the mechanisms causing cough in patients with PNDS are unclear, it is likely that secretions including inflammatory mediators penetrate to the larynx, and possibly to the trachea, and stimulate the afferent limb of the cough reflex in the upper respiratory tract [133]. C. Laryngospasm
Exaggeration of upper airway defensive reflexes is also a clinical problem. An example of exaggerated reflex responses is laryngospasm. Laryngospasm usually does not occur during wakefulness but it frequently occurs during a light depth of anesthesia, suggesting that light anesthesia may potentiate or consciousness may attenuate the upper airway reflexes. Laryngospasm consists of prolonged intense laryngeal closure in response to direct laryngeal stimulation from inhaled irritant agents, secretions or foreign bodies, leading to severe hypoxemia. In both anesthetized animals and anesthetized humans, there is some evidence to suggest that the laryngeal responses to airway irritation interact with background chemical ventilatory drive. For example, in anesthetized cats hypercapnia alone and hypoxia alone decreases the degree and duration of laryngospasm due to SLN stimulation, whereas hypocapnia augments and prolongs the duration of this laryngospasm [134]. In addition, in humans the addition of 5% CO2 to inspired isoflurane significantly reduces the frequency and severity of airway complications such as breathholding, coughing, and laryngospasm [135]. D. Airway Obstruction
Recognition of clinical problems such as obstructive sleep apnea (OSA) has generated an immense interest in the patency of the upper airway in recent years. Upper airway collapse is often seen in anesthetized patients and in patients with OSA syndrome. Upper airway obstruction in unconscious subjects has been attributed primarily to a result of the tongue falling back [136], but more recent studies indicate that the most common site at which obstruction occurs is the pharynx [137,138]. Upper airway obstruction occurs easily in older, male, and overweight patients who frequently have reduced pharyngeal size or pharyngeal structural abnormalities, suggesting that anatomical factors play an important role in initiating the upper airway obstruction [139]. Upper airway patency during wakefulness is in large part attributable to continual control by the higher nervous system, which regulates inspiratory motor output to the muscles of the pharynx and related structures. In addition to
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the effects of upper airway dilating muscles, the thoracic muscles may also influence the upper airway. During inspiration with the diaphragm, a negative pressure is generated in the pharynx and this negative pressure during inspiration is considered an important factor in promoting occlusion of the upper airway. However, the negative-pressure reflex would be initiated during upper airway obstruction and would tend to compensate for airway obstruction while increasing the pharyngeal size or stiffness. In healthy subjects, nasal occlusion with large negative pressures does not collapse the upper airway, presumably because the collapsing effect of negative pressure is opposed by the action of airway dilating muscles. Moreover, in normal human subjects, topical anesthesia of the upper airway increases the incidence of episodes of airway obstruction during sleep, particularly if the pharynx is anesthetized [140,141]. Thus, at least in some circumstances, upper airway reflex responses may contribute to upper airway patency. Patients with OSA have small pharyngeal size and thus may have greater changes in upper airway pressure with ventilation, and hence reflex activation of the muscles of the airway. However, any pressure reflex may not be adequate to keep the upper airways patent in these patients during sleep. Patients with OSA may be at risk for hypoxemia not only during anesthesia but also after anesthesia and surgery. Although the effects of anesthesia and surgery on postoperative hypoxemia have not been fully examined in patients with OSA, extreme episodic hypoxemia has been reported in these patients [142,143]. The study of Isono et al. [144] showed that postoperative episodic hypoxemia is associated with the presence of preoperative sleep-disordered breathing. Although anesthesia and sedation often reduce both respiratory drive and airway reflex activity, the reduction in activity of upper airway muscles seems to be greater than that of the diaphragm [145]. Thus, it is possible that the imbalance between dilating and collapsing forces may be a precipitating factor of upper airway obstruction. The majority of awake patients are nasal breathers during quiet breathing. Thus, even in the presence of a patent upper airway, airway obstruction may occur if the ability to change from the nasal route of airflow to the oral route in response to nasal obstruction is impaired [146]. E.
Cardiovascular Responses
Mechanical or chemical stimulation of most parts of the respiratory tract causes hypertension as a primary reflex effect, suggesting that release of catecholamines from the adrenal medulla and/or sympathetic stimulation of the heart are involved in this primary reflex response [24]. However, quantitative evaluation of the circulatory response to airway irritation
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during spontaneous breathing is by no means easy, because the primary reflex changes in blood pressure may be masked by the secondary homodynamic effects of respiratory responses such as coughing, expiration reflex, and breath-holding, which cause considerable changes in intrathoracic pressures and thereby venous return. In lightly anesthetized, paralyzed humans, sudden increases in blood pressure and heart rate are frequently observed during laryngoscopy and tracheal intubation and these responses may be potential problems to patients with coronary artery disease [147]. The responses of hypertension and tachycardia may be diminished by pretreatment with topical anesthesia, intravenous lidocaine, narcotics, or b-adrenergic receptor blocking drugs; and ensuring an adequate depth of anesthesia at the time of airway stimulation [148]. VIII.
Conclusions
The majority of afferent nerves arising from the airways are conducted in the trigeminal nerve, vagal nerve, and their branches. The importance of these nerves’ afferent innervation of the airways and their roles in elicitation of airway reflexes can be demonstrated by selective blockade of these afferents. Although the types of airway reflexes vary with the site of stimulation, airway reflexes fall into two categories: (1) defensive or protective reflexes that protect the respiratory tract from potentially harmful influences, and (2) regulatory reflexes that determine the pattern of breathing, control smooth muscle tone, and maintain airway patency. In experimental studies, different types of receptors have been identified in the upper and lower airways and their properties have been clarified, based on single-fiber recordings. There are at least five different types of receptors in the upper airway and four different types of receptors in the lower airways. Although some of these receptors have been implicated in playing important roles in particular airway reflexes, it is rather difficult to say exactly which receptors are responsible for which reflexes. In other words, the link between a given receptor and a particular reflex is unclear. Concerning airway receptors and airway reflexes, most information has been obtained in anesthetized animals and relatively few studies have evaluated the role of airway reflexes in humans. The application of experimental studies to pathophysiology in human subjects and patients would be significant, although it remains to be determined whether the results obtained from animal studies may be extended to humans. The better understanding of airway reflexes is essential to the better management of patients who need respiratory care.
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Acknowledgments This work was supported in part by a grant for the Second-term Comprehensive 10 years Strategy for Cancer Control from the Ministry of Health, Labour and Welfare of Japan. I am grateful to Dr. Franca B. Sant’Ambrogio for her critical reading of this chapter. References 1.
2.
3.
4. 5.
6. 7. 8.
9.
10.
11. 12.
13.
14.
Dietrich, W.D., Lowry, O.H. and Loewy, A.D., The distribution of glutamate, GABA and aspartate in the nucleus tractus solitarius of the cats, Brain Res. 237, 254–260, 1982. Hwang, B. and Wu, J.-Y., Ultrastructural studies on catecholaminergic terminals and GABAergic neurons in nucleus solitarius of the rat medulla oblongata, Brain Res. 302, 57–67, 1984. Dashwood, M.R., Muddle, J.R. and Spyer, K.M., Opiate receptor subtypes in the nucleus tractus solitarii of the cat: The effect of vagal section, Eur. J. Pharmacol. 155, 85–91, 1988. Kamai, J., Role of opioidergic and serotonergic mechanism in cough and antitussives, Pulm. Pharmacol. 19, 349–356, 1996. Kamai, J., Hukuhara, T. and Kasuya, Y., Dopaminergic control of the cough reflex as demonstrated by the effects of apomorphine, Eur. J. Pharmacol. 141, 153–158, 1987. Kamai, J., Tanihara, H., Igarashi, H. and Kasuya, Y., Effects of N-methyl-Daspartate antagonists on the cough reflex, Eur. J. Pharmacol. 168, 152–158, 1989. Hatakeyama, S., Histological study of the nerve distribution in the larynx in the cat, Arch. Jap. Histol. 19, 369–389, 1960. Fillenz, M. and Widdicombe, J.C., Receptors of the lungs and airways, in Handbook of Sensory Physiology, Vol. 3, Neil, E., ed., Heidelberg, SpringerVerlag, pp. 81–112, 1971. Boushey, H.A., Richardson, P.S., Widdicombe, J.G. and Wise, J.C.M., The response of laryngeal afferent fibres to mechanical and chemical stimuli, J. Physiol. (Lond.) 240, 153–175, 1974. Sant’Ambrogio, G., Anderson, J.W., Sant’Ambrogio, F.B. and Mathew, O.P., Responses of laryngeal receptors to water solutions of different osmolality and ionic composition, Respir. Med. 85: (suppl A), 57–60, 1991. Sant’Ambrogio, G., Mathew, O.P., Sant’Ambrogio, F.B. and Fisher, J.T., Laryngeal cold receptors, Respir. Physiol. 59, 35–46, 1985. Sant’Ambrogio, G., Mathew, O.P., Fisher, J.T. and Sant’Ambrogio, F.B., Laryngeal receptors responding to transmural pressure, airflow and local muscle activity, Respir. Physiol. 54, 317–333, 1983. Glebovsky, V.D. and Bayev, A.V., Stimulation of nasal cavity mucosa trigeminal receptors with respiratory airflows, Sechenov. Physiol. J. USSR 70, 1534–1541, 1984. Wallois, F., Macron, J.M., Junieaux, V. and Duron, B., Trigeminal nasal receptors related to respiration and to various stimuli in cats, Respir. Physiol. 85, 111–125, 1991.
252
Nishino
15. Tsubone, H., Nasal ‘flow’ receptors of the rat, Respir. Physiol. 75, 51–64, 1989. 16. Sekizawa, S., Tsubone, H., Kuwahara, M. and Sugano, S., Nasal receptors responding to cold and L-menthol airflow in the guinea pig, Respir. Physiol. 103, 211–219, 1996. 17. Tusubone, H., Nasal ‘pressure’ receptors, Nippon Juigaku Zasshi 52, 225–232, 1990. 18. Agostoni, E., Chinnock, J.E., DeDaly, M.B. and Murray, J.G., Functional and histological studies of the vagus nerve and its branches to the heart, lungs and abdominal viscera in the cats, J. Physiol. (Lond.) 135, 182–205, 1957. 19. Jammes, Y., Nail, B., Mei, N. and Grimaud, C.H., Laryngeal afferents activated by phenyldiguanide and their response to cold air or helium–oxygen, Respir. Physiol. 67, 379–389, 1987. 20. Tsubone, H., Sant’Ambrogio, G., Anderson, J.W. and Orani, G.P., Laryngeal afferent activity and reflexes in the guinea pig, Respir. Physiol. 87, 157–164, 1992. 21. Palecek, F., Mathew, O.P., Sant’Ambrogio, F.B. and Sant’Ambrogio, G., Cardiorespiratory responses to inhaled laryngeal irritants, Inhal. Toxicol. 2, 93– 104, 1990. 22. Wallois, F., Gros, F., Masmoudi, K. and Larnicol, N., Trigeminal afferents implied in the triggering or inhibition of sneezing in cats, Neurosci. Lett. 122, 145–147, 1991. 23. Lundblad, L., Protective reflexes and vascular effects in the nasal mucosa elicited by activation of capsaicin-sensitive substance P-immunoreactive trigeminal neurons, Acta Physiol. Scand. (suppl) 529, 1–42, 1984. 24. Widdicombe, J.G., Reflexes from the upper respiratory tract, in Handbook of Physiology, Section 3 The Respiratory System, Vol. II, Part I, Cherniack, N.S. and Widdicombe, J.G., eds., Baltimore, American Physiological Society, Williams & Wilkins Co., pp. 363–394, 1986. 25. Wealthall, S.R., Factors resulting in failure to interrupt apnea, in Development of Upper Respiratory Anatomy and Function. Implications for Sudden Infant Death Syndrome, Bosma, J.P. and Showacre, J., eds., Washington, DC, US Government Printing Office, pp. 212–225, 1976. 26. Karlsson, J.-A., Sant’Ambrogio, G. and Widdicombe, J.G., Afferent neural pathways in cough and reflex bronchoconstriction, J. Appl. Physiol. 65, 1007– 1023, 1988. 27. Szereda-Przestaszewska, M. and Widdicombe, J.G., Reflex effects of chemical irritation of the upper airways on the laryngeal lumen in cats, Respir. Physiol. 18, 107–115, 1973. 28. Hinkel, J.E. and Tantum, K.R., A technique of measuring reactivity of the glottis, Anesthesiology 35, 634–637, 1971. 29. Korpas, J., The expiration reflex from vocal cords, Physiol. Bohemoslov. 21, 408–409, 1972. 30. Korpas, J. and Tomori, S., Cough and Other Respiratory Reflexes, Larger, Basel, pp. 15–188, 1979. 31. Stockwell, M., Lang, S., Yip, T., Zintel, T., White, C. and Gallagher, C.G., Lack of importance of the superior laryngeal nerves in citric acid cough in humans, J. Appl. Physiol. 75, 613–617, 1993.
Airway Reflexes in Humans 32.
33.
34. 35. 36.
37. 38.
39.
40.
41.
42.
43. 44. 45.
46.
47. 48.
49.
253
Higenbottam, T., Jackson, M., Woolman. P., Lowry, R. and Wallwork, J., The cough response to ultrasonically nebulized distilled water in heart–lung transplantation patients, Am. Rev. Respir. Dis 140, 58–61, 1989. Tomori, Z. and Widdicombe, J.G., Muscular, bronchomotor and cardiovascular reflexes elicited by mechanical stimulation of the respiratory tract, J. Physiol. (Lond.) 200, 25–50, 1969. Nishino, T., Swallowing as a protective reflex for the upper respiratory tract, Anesthesiology 79, 588–601, 1993. Miller, A.J., Deglutition, Physiol. Rev., 62, 129–184, 1982. Nishino, T., Yonezawa, T. and Honda, Y., Effects of swallowing on the pattern of continuous respiration in human adults, Am. Rev. Respir. Dis 132, 1319– 1322, 1985. Nishino, T. and Hiraga, K., Coordination of swallowing and respiration in unconscious subjects, J. Appl. Physiol. 70, 988–993, 1991. Nishino, T., Hasegawa, R., Ide, T. and Isono, S., Hypercapnia enhances the development of coughing during continuous infusion of water into the pharynx, Am. J. Respir. Care Med. 157, 815–821, 1998. Kijima, M., Isono, S. and Nishino, T., Coordination of swallowing and phase of respiration during added respiratory loads in awake subjects, Am. Respir. Crit. Care Med. 159, 1898–1902, 1999. Paydafar, D., Gilert, R.J., Poppel, C.S. and Nassabb, P.F., Respiratory phase resetting and airflow changes induced by swallowing in humans, J. Physiol. (Lond.) 483, 273–288, 1995. Hammouda, M. and Wilson, W.H., Influences which affect the form of the respiratory cycle, in particular that of the expiratory phase, J. Physiol. (Lond.) 80, 261–284, 1933. Mathew, O.P., Abu-Osba, Y.K. and Thach, B.T., Influence of upper airway pressure changes on genioglossus muscle respiratory activity, J. Appl. Physiol. 52, 438–444, 1982. Mathew, O.P., Abu-Osba, Y.K. and Thach, B.T., Influence of upper airway pressure changes on respiratory frequency, Respir. Physiol. 49, 223–233, 1982. Mathew, O.P., Upper airway negative-pressure effects on respiratory activity of upper airway muscles, J. Appl. Physiol. 56, 500–505, 1984. Roberts, J.L., Reed, W.R., Mathew, O.P., Menon, A.A. and Thach, B.T., Assessment of pharyngeal airway stability in normal and micrognathic infants, J. Appl. Physiol. 58, 290–299, 1985. Carlo, W.A., Miller, M.J. and Martin, R.J., Differential responses of respiratory muscles to airway occlusion in infants, J. Appl. Physiol. 59, 847– 852, 1985. Van Lunteren, E. and Strohl, K.P., The muscles of the upper airways, Clin. Chest Med. 7, 171–188, 1986. Horner, R.L., Innes, J.A., Murphy, K. and Guz, A., Evidence for reflex upper airway dilator muscle activation by sudden negative airway pressure in man, J. Physiol. 436, 15–29, 1991. Van Lunteren, E., Strohl, K.P., Parker, D.M., Bruce, E.N., Van de Graff, W.B. and Cherniack, N.S., Phasic volume-related feedback on upper airway muscle activity, J. Appl. Physiol. 56, 730–736, 1984.
254 50.
51. 52.
53.
54.
55. 56. 57. 58.
59. 60.
61.
62.
63. 64.
65. 66.
67.
Nishino Liistro, G., Stanescu, D.C., Veriter, C., Rodenstein, D.O. and D’Odemont, J.P., Upper airway anesthesia induces airflow limitation in awake humans, Am. Rev. Respir. Dis. 146, 581–585, 1992. Leiter, J.C. and Daubenspeck, J.A., Selective reflex activation of the genioglossus in humans, J. Appl. Physiol. 68, 2581–2587, 1990. Horner, R.L., Innes, J.A., Holden, H.B. and Guz, A., Afferent pathway(s) for pharyngeal dilator reflex to negative airway pressure in man: A study using upper airway anesthesia, J. Physiol. (Lond.) 436, 31–44, 1991. Horner, R.L., Innes, J.A., Morrel, M.J., Shea, S.A. and Guz, A., The effect of sleep on reflex genioglossus muscle activation by stimuli of negative airway pressure in humans, J. Physiol. (Lond.) 476, 141–151, 1994. Thach, B.T., Schefft, G.L., Pickens, D.L. and Menon, A.P., Influence of upper airway negative pressure reflex on response to airway occlusion in sleeping infants, J. Appl. Physiol. 67, 749–755, 1989. Nishino, T. and Kochi, T., Effects of sedation produced by thiopentone on responses to nasal occlusion in female adults, Br. J. Anaesth. 71, 388–392, 1993. Nishino, T., Tanaka, A. and Ishikawa, T., Effects of topical nasal anaesthesia on shift of breathing route in adults, Lancet 339, 1497–1500, 1992. Richardson, J. and Beland, J., Non-adrenergic inhibitory nervous system in human airways, J. Appl. Physiol. 41, 764–771, 1976. Partanen, M., Laitinen, A., Hervonen, A., Toivanen, M. and Laitinen, L.A., Catecholamine- and acetylcholinesterase-containing nerves in human lower respiratory tract, Histochemistry 76, 175–188, 1982. Widdicombe, J.G., Airway receptors, Respir. Physiol. 125, 3–15, 2001. Nishino, T., Anderson, J.W. and Sant’Ambrogio, G., Responses of tracheobronchial receptors to halothane, enflurane, and isoflurane in anesthetized dogs, Respir. Physiol. 95, 281–294, 1994. Matsumoto, S., Nagayama, T., Yamasaki, M., Kanno, T. and Shimizu, T., Cholinergic and H1-receptor influences of histamine on slowly adapting pulmonary stretch receptor activity in the rabbit, J. Auton. Nerv. Syst. 40, 107– 120, 1992. Matsumoto, S., Effects of vagal stimulation on slowly adapting pulmonary stretch receptors and lung mechanics in anesthetized rabbits, Lung 174, 333–344, 1996. Sant’Ambrogio, G., Information arising from the tracheobronchial tree of mammals, Physiol. Rev. 62, 531–569, 1982. Matsumoto, S., Takahashi, T., Tanimoto, T., Saiki, C. and Takeda, M., Effects of potassium channel blockers on CO2-induced slowly adapting pulmonary stretch receptor inhibition, J. Pharmacol. Exp. Ther. 290, 974–979, 1999. Widdicombe, J.G., Neurophysiology of the cough reflex, Eur. Respir. J. 8, 1193–1202, 1995. Mutoh, T., Tsubone, H., Nishimura, R. and Sasaki, N., Effects of volatile anesthetics on vagal C-fiber activities and their reflexes in anesthetized dog, Respir. Physiol. 112, 253–264, 1998. Kaufman, M.P., Iwamoto, G.A. Ashton, J.H. and Cassidy, S.S., Responses to inflation of vagal afferents with endings in the lungs of dog, Circ. Res. 51, 525–531, 1982.
Airway Reflexes in Humans 68.
69.
70.
71.
72.
73.
74.
75. 76. 77. 78. 79. 80. 81. 82. 83.
84. 85. 86.
255
Coleridge, H.M. and Coleridge, J.C.G., Afferent vagal C fibre innervation of the lungs and airways and its functional significance, Rev. Physiol. Biochem. Pharmacol. 99, 1–110, 1984. Coleridge, H.M. and Coleridge, J.C.G., Reflexes evoked from tracheobronchial tree and lungs, in Handbook of Physiology, Section 3, The Respiratory System, Vol. II, Part I, Cherniack, N.S. and Widdicombe, J.G., eds., Baltimore, American Physiological Society, pp. 395–429, 1986. Brouns, I., Van Genechten, J., Hayashi, H., Gajda, M., Gomi, T., Burnstock, G., Timmermans, J.P. and Adriaensen, D., Dual sensory innervation of pulmonary neuroepithelial bodies, Am. J. Respir. Cell Mol. Biol. 28, 275–285, 2003. Gautier, H., Bonora, M. and Gaudy, J.R., Breuer-Hering inflation reflex and breathing pattern in anesthetized humans and cats, J. Appl. Physiol. 51, 1162–1168, 1981. O’Kelly, I., Stephens, R.H., Peers, C. and Kemp, P.J., Potential identification of the O2-sensitive Kþ current in a human neuroepithelial body-derived cell line, Am. J. Physiol. 276, L96–L104, 1999. Tryfon, S., Kontakiotis, T., Mavrofridis, E. and Patakas, D., Hering-Breuer reflex in normal adults and in patients with chronic obstructive pulmonary disease and interstitial fibrosis, Respiration 68, 140–144, 2001. Hamilton, R.D., Winning, A.J., Horner, R.L. and Guz, A., The effect of lung inflation on breathing in man during wakefulness and sleep, Respir. Physiol. 73, 145–154, 1988. Guz, A., Noble, M.I., Eisele, J.H. and Trenchard, D., The effect of lung deflation on breathing in man, Clin. Sci. 40, 451–461, 1971. Koller, E.A. and Ferrer, P., Studies on the role of the lung deflation reflex, Respir. Physiol. 10, 172–183, 1970. Green, J.F. and Kaufman, M.P., Pulmonary afferent control of breathing as end-expiratory lung volume decreases, J. Appl. Physiol. 68, 2186–2194, 1990. Widdicombe, J.G. and Nadel, J.A., Reflex effects of lung inflation on tracheal volume, J. Appl. Physiol. 18, 681–686, 1963. Widdicombe, J.G. and Nadel, J.A., Airway volume, airway resistance, and work and force of breathing: Theory, J. Appl. Physiol. 18, 863–868, 1963. Bucher K, Pathophysiology and pharmacology of cough, Pharmacol. Rev. 10, 43–58, 1958. Hanacek, J.A., Davies, J.A. and Widdicombe, J.G., Influence of lung stretch receptor on the cough reflex in rabbits, Respiration 45, 161–168, 1984. Sant’Ambrogio, G., Sant’Ambrogio, F.B. and Davies, A., Airway receptors in cough, Bull. Eur. Physiopathol. Respir. 20, 43–47, 1984. Nishino, T., Hiraga, Y. and Honda, Y., Influence of CPAP on reflex responses to tracheal irritation in anesthetized humans, J. Appl. Physiol. 67, 954–958, 1989. Korpas, J. and Kalosayova, G., The expiration reflex in mice, Physiol. Bohemoslov. 24, 253–256, 1975. Korpas, J. and Widdicombe, J.G., Defensive respiratory reflexes in ferrets, Respiration 44, 128–135, 1983. Widdicombe, J.G., Respiratory reflexes from the trachea and bronchi of the cat, J. Physiol. (Lond.) 123, 71–104, 1964.
256
Nishino
87.
Tomori, Z., Lemakova, S. and Hlecyova, A., Defensive reflexes of the respiratory tract in dogs, Physiol. Bohemoslov. 26, 49–54, 1977. Sullivan, C.E., Kozar, L., Murphy, E. and Phillipson, E.A., Arousal, ventilatory, and airway responses to bronchopulmonary stimulation in sleeping dogs, J. Appl. Physiol. 47, 17–25, 1979. Nishino, T. and Honda, Y., Respiratory reflex responses to stimulation of tracheal mucosa in enflurane-anesthetized humans, J. Appl. Physiol. 65, 1069–1074, 1988. Nishino, T., Tagaito, Y. and Isono, S., Cough and other reflexes on irritation of airway mucosa in man, Pulm. Pharmacol. 9, 285–292, 1996. Jackson, C., Cough: Bronchoscopic observations on the cough reflex, JAMA 79, 1399–1403, 1922. Lee, L.-Y., Kou, Y.R., Frazier, D.T., Beck, E.R., Pisarri, T.E. and Coleridge, J.C.G., Stimulation of vagal pulmonary C-fibers by a single breath of cigarette smoke in dogs, J. Appl. Physiol. 66, 2032–2038, 1989. Lee, L.-Y., Gerhardstein, D.C., Wang, A.L. and Burki, N.K., Nicotine is responsible for airway irritation evoked by inhaling cigarette smoke in man, J. Appl. Physiol. 75, 1955–1961, 1993. Schelegle, E.S., Carl, M.L., Coleridge, H.M., Coleridge, J.C.G. and Green, J.G., Contribution of vagal afferents to respiratory reflexes evoked by acute inhalation of ozone in dog, J. Appl. Physiol. 74, 2338–2344, 1993. Beckett, W.S., McDonnell, W.F., Horstman, D.H. and House, D.E., Role of the parasympathetic nervous system in acute lung response to ozone, J. Appl. Physiol. 59, 1879–1885, 1985. Hazucha, M.J., Bates, D.V. and Bromberg, P.A., Mechanism of action of ozone on human lung, J. Appl. Physiol. 67, 1535–1541, 1989. Coleridge, J.C.G., Coleridge, H.M., Schelegle, E.S. and Green, J.F., Acute inhalation of ozone stimulates bronchial C-fibers and rapidly adapting receptors in dogs, J. Appl. Physiol. 74, 2345–2352, 1993. Ho, C.Y. and Lee, L.-Y., Ozone enhances excitabilities of pulmonary C fibers to chemical and mechanical stimuli in anesthetized rats, J. Appl. Physiol. 85, 1509–1515, 1998. Fuller, R., The human pharmacology of capsaicin, Arch. Int. Pharmacodyn. Ther. 303, 147–156, 1990. Karlsson, J.A., The role of capsaicin-sensitive C-fibre afferent nerves in the cough reflex, Pulm. Pharmacol. 9, 315–321, 1996. Sekizawa, K., Jia, Y.X., Ebihara, T., Hirose, Y., Hirayama, Y. and Sasaki, H., Role of substance P in cough, Pulm. Pharmacol. 9, 323–328, 1996. Joos, G.F., Pauwels, R.A. and van der Straeten, M.E., Effect of inhaled substance P and neurokinin A on the airways of normal and asthmatic subjects, Thorax 42, 779–783, 1987. Katsumata, U., Sekizawa, K., Inoue, H., Sasaki, H. and Takishima, T., Inhibitory actions of procaterol, a beta-2 stimulant, on substance P-induced cough in normal subjects during upper respiratory tract infection, Tohoku J. Exp. Med. 158, 105–106, 1989. Winning, A.F., Hamilton, R.D., Shea, S.A. and Guz, A., Respiratory and cardiovascular effects of central and peripheral intravenous injections of
88.
89.
90. 91. 92.
93.
94.
95.
96. 97.
98.
99. 100. 101. 102.
103.
104.
Airway Reflexes in Humans
105.
106.
107. 108.
109.
110.
111.
112.
113.
114.
115.
116.
117. 118. 119.
120.
257
capsaicin in man: Evidence for pulmonary chemosensitivity, Clin. Sci. 71, 519–526, 1986. Tatar, M., Webber, S.E. and Widdicombe, J.G., Lung C-fibre receptor activation and defensive reflexes in anaesthetized cats, J. Physiol. (Lond.) 402, 411–420, 1988. Mohammed, S.P., Higenbottam, T.W. and Adcock, J.J., Effects of aerosolapplied capsaicin, histamine and prostaglandin E2 on airway sensory receptors of anaesthetized cats, J. Physiol. (Lond.) 469, 51–66, 1993. Marik, P.E., Aspiration pneumonitis and aspiration pneumonia, N. Eng. J. Med. 344, 665–671, 2001. Roy, T.M., Ossorio, M.A., Ciipolla, L.M., Fields, C.L., Snider, H.L. and Anderson, W.H., Pulmonary complications after tricyclic antidepressant overdose, Chest 96, 852–856, 1989. Tiret, L., Nivoche, Y., Hatton, F., Desmonts, J.M. and Vourc’h, G., Complications related to anaesthesia in infants and children. A prospective survey of 40240 anaesthetics, Br. J. Anaesth. 61, 263–269, 1988. Olsson, G.L., Hallen, B. and Hambraeus-Jonzon, K., Aspiration during anaesthesia: A computer-aided study of 185,358 anaesthetics, Acta Anaesthesiol. Scand. 30, 84–92, 1986. Marrie, T.J., Durant, H. and Yates, L., Community-acquired pneumonia requiring hospitalization: 5-year prospective study, Rev. Infect. Dis. 11, 586– 599, 1989. Torres, A., Serra-Batlles, J., Ferrer, A., Jimenez, P., Celis, R., Cobo, E. and Rodriguez-Roisin, R., Severe community-acquired pneumonia: Epidemiology and prognostic factors, Am. Rev. Respir. Dis. 144, 312–318, 1991. Moine, P., Vercken, J.P., Chevret, S., Chatang, C. and Gajdos, P., Severe community-acquired pneumonia: Etiology, epidemiology, and prognosis factors, Chest 105, 1487–1495, 1994. Gotta, A.W. and Sullivan, C.A., Anaesthesia of the upper airway using a topical anaesthetic and superior laryngeal nerve block, Br. J. Anaesth. 53, 1055–1058, 1981. Brock-Utne, J.G., Winning, T.J., Rubin, J. and Kingston, H.G.G., Laryngeal incompetence during neuroleptanalgesia in combination with diazepam, Br. J. Anaesth. 48, 699–701, 1976. Rubin, J., Brock-Utne, J.G., Greenberg, M., Bortz, J. and Dowing, J.W., Laryngeal incompetence during experimental ‘relative analgesia’ using 50% nitrous oxide in oxygen, Br. J. Anaesth. 49, 1005–1008, 1977. Brin, M.F. and Younger, D., Neurologic disorders and aspiration, Otolaryngol. Clin. North Am. 21, 691–699, 1988. Irwin, R.S. and Curley, F.J., The treatment of cough: A comprehensive review, Chest 6, 1477–1484, 1991. Irwin, T.S., Boulet, L.-P., Coutier, M.M., Gold, R.M., Ing, A.J., O’Byrne, P., Prakash, U.B.S., Pratter, M.R. and Rubin, B.K., Managing cough as a defense mechanism and as a symptom: A consensus panel report of the American College of Chest Physicians, Chest Suppl, 133S–181S, 1998. Stone, R., Chronic cough: Mechanisms and management, Respir. Med. 87, 249–251, 1993.
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Nishino
121. Baitner, A.C., Bernstein, A.D., Jazrawi, A.J., Della Valle, C.J. and Jazrawi, L.M., Spontaneous rib fracture during pregnancy. A case report and review of the literature, Bull. Hosp. Jt. Dis. 59, 163–165, 2000. 122. Sharpey-Shafer, E.P., The mechanism of syncope after cough, Fr. Med. J. 2, 860–863, 1953. 123. Haider, A.W., Larson, M.G., O’Donnell, C.J., Evans, J.C., Wilson, P.W. and Levy, D., The association of chronic cough with the risk of myocardial infarction: The Framingham Heart Study, Am. J. Med. 106, 279–284, 1999. 124. Irwin, R.S., Curley, F.G. and French, C.L., Chronic cough: The spectrum and frequency of causes, key components of the diagnostic evaluation, and outcome of specific therapy, Am. Rev. Respir. Dis. 141, 640–647, 1990. 125. Choudry, N.B. and Fuller, R.W., Sensitivity of the cough reflex in patients with chronic cough, Eur. Respir. J. 5, 296–300, 1992. 126. Berkin, K.E., Respiratory effects of angiotensin converting enzyme inhibition, Eur. Respir. J. 2, 198–201, 1989. 127. Morice, A.H., Lowry, R., Brown, N.J. and Higenbottam, T., Angiotensinconverting enzyme and the cough reflex, Lancet 2, 1116–1118, 1987. 128. Irwin, R.S., Zawaki, X., Curley, F.J., French, C.L. and Hoffman, P.J., Chronic cough as the sole presenting manifestation of gastroesophageal reflux, Am. Rev. Respir. Dis. 140, 1294–1300, 1989. 129. Ing, A.J., Ngu, M.C. and Breslin, A.B.X., Pathogenesis of chronic persistent cough associated with gastroesophageal reflux, Am. J. Respir. Crit. Care Med. 149, 160–167, 1994. 130. Carrao, W.M., Braman, S.S. and Irwin, R.S., Chronic cough as the sole presenting manifestation of bronchial asthma, N. Engl. J. Med. 300, 633–637, 1979. 131. O’Connell, E.J., Rojas, A.R. and Sachs, M.I., Cough-type asthma: A review, Ann. Allergy 66, 279–282, 1991. 132. Johnson, D. and Osborn, L.M., Cough variant asthma: A review of the clinical literature, J. Asthma 28, 85–90, 1991. 133. Irwin, R.S., Pratter, M.R., Holland, P.S., Corwin, R.W. and Hughes, J.P., Postnasal drip causes cough and is associated with reversible upper airway obstruction, Chest 85, 346–352, 1984. 134. Nishino, T., Yonezawa, T. and Honda, Y., Modification of laryngospasm in response to changes in PaCO2 and PaO2 in the cat, Anesthesiology 55, 286– 291, 1981. 135. Coleman, S.M., McCrory, J.W., Vallis, C.J. and Boys, R.J., Inhalation induction of anesthesia with isoflurane: Effect of added carbon dioxide, Br. J. Anaesth. 67, 257–261, 1991. 136. Safar, P., Escarraga, L.A. and Chang, F., Upper airway obstruction in the unconscious patient, J. Appl. Physiol. 14, 760–764, 1959. 137. Nandi, P.R., Charlesworth, C.H., Taylor, S.J., Nunn, J.F. and Dore, C.J., Effect of general anaesthesia on the pharynx, Br. J. Anaesth. 66, 157–162, 1991. 138. Hudgel, D.W. and Hendricks, C., Palate and hypopharynx: Site of inspiratory narrowing of the upper airway during sleep, Am. Rev. Respir. Dis. 138, 1542–1547, 1988.
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139. Isono, S., Remmers, J.E., Tanaka, A., Sho, Y. and Nishino, T., Static properties of the passive pharynx in sleep apnea, Sleep 19, S175–S177, 1996. 140. McNicholas, W.T., Coffey, M., McDonnell, T., O’Regan, R. and Fitzgerald, M.X., Upper airway obstruction during sleep in normal subjects after selective topical oropharyngeal anesthesia, Am. Rev. Respir. Dis. 135, 1316–1319, 1987. 141. Fogel, R.B., Malhotra, A., Shea, S.A., Edwards, J.K. and White, D.P., Reduced genioglossal activity with upper airway anesthesia in awake patients with OSA, J. Appl. Physiol. 88, 1346–1354, 2000. 142. Rosenberg, J. and Kehlet, H., Postoperative episodic oxygen desaturation in the sleep apnea syndrome, Acta Anaesth. Scand. 35, 368–369, 1991. 143. Reeder, M.K., Goldman, M.D., Loh, L., Muir, A.D., Casey, K.R. and Lehance, J.R., Late postoperative nocturnal dips in oxygen saturation in patients undergoing major abdominal surgery, Anesthesia 47, 110–115, 1992. 144. Isono, S., Sha, M., Suzukawa, M., Sho, Y., Ohmura, A., Kudo, Y., Misawa, K., Inaba, S. and Nishino, T., Preoperative nocturnal desaturations as a risk factor for late postoperative nocturnal desaturations, Br. J. Anaesth. 80, 602– 605, 1998. 145. Nishino, T., Honda, Y., Kohchi, T., Shirahata, M. and Yonezawa, T., Comparison of changes in the hypoglossal and phrenic nerve activity in response to increasing depth of anesthesia in cats, Anesthesiology 60, 19–24, 1984. 146. Cook, T.A. and Komorn, R.M., Statistical analysis of the alterations of blood gases produced by nasal packing, Laryngoscope 83, 1802–1809, 1973. 147. Forbes, A.M. and Dally, F.G., Acute hypertension during induction of anesthesia and endotracheal intubation in normotensive man, Br. J. Anaesth. 42, 618–624, 1970. 148. Donlon, J.V., Jr., Anesthesia and eye, ear, nose, and throat surgery, in Anesthesia, 4th edn., Vol. II, Miller, R.D., ed., New York, Churchill Livingstone, pp. 2175–2196, 1994.
8 Inheritance and Ventilatory Behavior in Animal Models
KINGMAN P. STROHL Case Western Reserve University Cleveland, Ohio
This review concerns the genetic, non-environmental factors that influence normal variations in ventilation and/or its components, frequency and tidal volume. The question of ‘is there a genetic effect?’ is now answered a number of times in both human studies and rodent models, and there is moderate evidence to suspect that specific gene regions operate in determining the apnea–hypopnea index, a defining value for human sleep apnea. There is the best evidence in animal models to the address questions of ‘how strong is the genetic component?’ and ‘what genes might be involved?’ There is consensus regarding the collection of phenotype values for frequency and tidal volume and the response to chemosensory challenges, and qualitative and quantitative differences exist among rodent strains in both steady-state and transient changes in these traits to chemosensory challenge. Some gene regions could be interesting in regard to explaining the risk of progression or of severity of diseases in which disorders of ventilatory control operate to produce hypoxic complications. Computational analyses of datasets and breeding of animals offer 261
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opportunity to create a functional map of the connections between genes and ventilatory behavior. I.
Introduction
Ventilatory rate and depth (ventilatory behavior) alters its characteristics during growth and development as well as in response to both short- and long-term perturbations to preserve homeostasis in cellular respiration, acid–base balance, and heat exchange. In mammals, ventilation is achieved through combinations of tidal volume and frequency [1], yet there exists a wide range of respiratory frequencies that are mechanically efficient [2,3]. This flexibility allows ample scope for genetic factors to modify ventilatory behavior in a homeostatic manner without an increase in energy cost. Diseases of the chest wall and lungs restrict this range, produce a mismatch between respiratory control and ventilatory effects, and are associated with feelings of breathlessness or dyspnea [4,5]. In sleep-disordered breathing, the problem in ventilatory control is state-related; disturbances in ventilation fragment sleep and produce hypoxic stress resulting in daytime symptoms of sleepiness and cardiovascular sequelae [6]. This chapter will review the evidence that reproducible differences in ventilatory behavior occur between otherwise healthy animals. While individual values are modified over time by factors such as chronic exposure to altitude, respiratory loading, physical fitness, age, hormones, sex hormones, and sleep [7,8], the focus of this review will be the inheritance of ventilatory traits in animal models (mice and rats). Such models can yield insight into clinical disorders of respiratory control, such as sleep apnea. II.
Evidence and Implications for Inheritance of Ventilatory Traits in Humans
One can identify an individuality in respiratory patterning of tidal volume and frequency in humans during both wakefulness and sleep [9]. The shape and timing of tidal breaths are more similar in monozygotic twins than in unrelated individuals, not only during breathing at rest but also during behavioral tasks as well [10]. These and other observations suggest that variations in respiratory patterning occur in healthy humans and may have an inherited basis. Furthermore, such variation is not merely the result of differences in mechanical properties or in respiratory gas exchange. Thus, the causes for this phenomenon reside in central patterning of respiratory rate and the size and pattern of tidal volume [11]. Additional evidence indicates that humans exhibit heritable variations in the response to standard ventilatory challenges. There are absent or weak hypercapnic responses in natives of Papua New Guinea in contrast to the
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two- to four-fold increases in ventilatory responsiveness to carbon dioxide present in residents of Australia [12]. Sibling similarity occurs in the blunted ventilatory responses to hypercapnia first reported in athletes [13]. Others suggest familial clustering of hypoxic sensitivity is a distinguishing characteristic in this same population [14]. There are greater similarities in hypoxic ventilatory responsiveness between monozygotic (MZ) twins than between dyzygotic (DZ) twins at sea level [15–19]. Some but not all studies reported concordance of patterns of ventilatory responsiveness to hypercapnia between MZ twins [20,21]. Reports indicate abnormal ventilatory responsiveness to hypercapnia or hypoxia is present in first-degree relatives of patients with excessive hypercapnia, hypoxia, or unexplained respiratory failure [20,22–25] or those with sleep apnea [26]. This literature forms one basis for concluding that there occurs a rather wide range of variation in chemoresponsiveness in the healthy human population, and that such variation is to some significant degree likely to result from inheritable factors. The literature on familial factors that operate in the presentation of sleep apnea is rather good [27]. While direct studies of genetic factors in the cause of clinical disorders are limited in number, there is more direct evidence in assignment of genetic strength. There is an increased prevalence (two-fold) in a polymorphism for apolipoprotein E in patients with sleep apnea [28], a polymorphism also associated with cardiovascular disease and Alzheimer’s disease. This is not observed in other studies [29,30], possibly because of ascertainment bias. In the Cleveland cohort, there is statistical evidence for an oligogenic transmission explaining some 27% of the variation in apnea–hypopnea index (AHI) expression in the community. Having two or more family members with elevated values for AHI is associated with increased likelihood of finding another (Figure 8.1) [31]. In addition, families in Cleveland with a history consistent with sudden infant death more often have two or more members with obstructive sleep apnea (OSA), when compared with families without such history. Other associations in this dataset include a tendency for a more blunted hypoxic response, and cephalometric measures of brachycephaly, a smaller mean posterior nasal spine–basion distance (smaller posterior airway), and smaller mandibular/maxillary ratio (shorter jaw) [32]. Thus there are interesting components in the respiratory control system that appear to infer inheritable risk for complex process like sleep apnea. Even more direct is the evidence from a 10 cM genome scan on some members of the Cleveland Family Study [33]. Individuals were selected to be maximally informative for AHI, a defining metric for obstructive sleep apnea. Subjects were genotyped for 375 autosomal microsatellite markers at a sex-averaged spacing of 9.1 cM (Marshfield Center for Human Genetics, set 10). The sample included 66 extended pedigrees of European–American origin comprising 344 subjects and 59 African–American pedigrees (n ¼ 257
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2
1.5
>3
1 0.5 0 1
2
>3
Number of family members with sleep apnea
Figure 8.1 Shown are the odds ratios for the probability of finding another person with an elevated apnea–hypopnea index (AHI) based upon the number of people in a family currently identified with an elevated AHI. There are instances where it appears that sleep apnea has a strong familial component (Data from Ref. 27).
individuals). A multipoint model-free linkage analysis of the logtransformed AHI and body mass index (BMI) was performed (modelbased analyses also were performed). In order to utilize all of the available phenotypic and genotypic information, a variance components approach appropriate for extended pedigrees was used [33]. A summary of these findings is presented in Table 8.1. The linkage scores are modest; however, the sensitivity of the scan is also only modest, being model-free with an 10 cM average length. As well, there is the possibility of heterotypes, with pedigrees in which different gene regions contributing to the expression of AHI and/or BMI phenotype. One area of linkage was to a marker on chromosome 19 that resides in the region of the apolipoprotein E (APOE) locus, a region already of interest in sleep apnea [28]. A second biologically plausible and clinically interesting candidate gene, proopiomelanocortin (POMC), resides in the region of maximal linkage on chromosome 2. The sleep apnea phenotype and the POMC linkage are consistent with the proposed relationship of sleep-disordered breathing to the metabolic syndrome [34]. Finding such linkage allows one to consider studies on the separate and combined effects of obesity and diet in rodent models. There is an interplay between melanocortin and neuropeptide Y (NPY) in pathological weight loss and weight gain in murine models [35]. POMC is of central importance to energy metabolism,
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Table 8.1 Gene Regions in the Human Associated with Sleep Apnea Trait
Human chromosome
Distance (cM)
LOD scorea
AHI
2 19 8c 19 8c 2 12
74b
1.4–1.6
65 137 54 72
1.2 1.5 1.8 1.7
BMI a
Adjusted for age and gender. Region-of-interest includes proopiomelanocortin (POMC). c A–A only. Source: Data from Ref. 182. b
leptin levels, and other features of the metabolic syndrome [36]. The human POMC gene is homologous to mouse Ch. 14 (4 cM), where polymorphisms among mouse strains are linked with the metabolic syndrome. Indeed, leptin resistance differs between strains of mice without genetically determined obesity at this linkage site, the A/J mouse being resistant and the C57BL/6J being sensitive to diet-induced obesity [37], and there exist differences between strains at a level of POMC expression [38]. Yet leptin supplementation in the C57BL/6J animals that show low plasma leptin levels in response to fat feeding slows but does not prevent the subsequent development of diet-induced obesity [39], suggesting a polygenic influence on trait expression. There may be gene polymorphisms that are disease-modifying rather than disease-causing. One gene of interest is the angiotensin-converting enzyme gene (ACE, human Chromosome 27). There are now two reports that link ACE I/D polymorphisms to apnea length and sleep hypoxemia in OSA patients [40,41]. In one European study, the frequency distribution of the I/D polymorphism was similar in the control population as in OSA patients; also, the levels of ACE were higher in OSA patients and fell with treatment [42], providing some support for ACE as a disease-modifying allele. Another set of candidate genes relates to the nitric oxide synthase (NOS) enzymes that produce NO (nitric oxide). There is some data that NO production is reduced in OSA patients [43,44], and that levels rise with treatment [45]. Yet one study suggests that NO levels increase from before sleep to after sleep in patients with sleep apnea [46]. This is an interesting result since at sea level acute and sub-acute exposures to hypoxia reduce NOS activity and NO production; moreover, high altitude natives at 4000 m often have higher levels of exhaled NO than sea level natives [47].
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Presumably a downregulation in the NO system in sleep apnea could be a factor in the altered vascular reactivity seen in this disorder [48], and be modeled by intermittent hypoxia [49]. A theme in all these studies may be that gene polymorphisms affect the quality, quantity, or consequences of abnormal breathing events during sleep. Therefore, there is reason to believe that efforts to quantify and identify the chromosomal mechanisms of sleep apnea may unravel the physiology of normal breathing and of hyper- or hypo-ventilation states. Studies of genetic factors complement studies of cellular and organ system models, and precedent exists for identifying homologues in the rat, mouse, and human genome [50,51]. Genetic homologies among species also provide an extraordinary opportunity for understanding the evolution of ventilation.
III.
Targeting Ventilatory Traits in Small Animals
Ventilatory behavior is one part of a respiratory control system that operates to provide sufficient oxygen to meet cellular metabolic requirements and to remove enough carbon dioxide so that cell function is not impaired by excessive change in hydrogen ion concentration. What is remarkable is that in the healthy state, the respiratory control system will maintain arterial PO2 and PCO2 within a fairly narrow range despite changes in metabolism and environmental conditions with growth, development, and extremes of daily living. Respiratory control operates using principles of feedback and feedforward control [7]. Figure 8.2 illustrates essential components of this engineered system. The controller regulates ventilatory behavior (the controlled system) through efferent neural pathways coordinating contraction and relaxation of the controlled variable (respiratory muscles). A multiplicity of inputs to the respiratory neurons ensures that ventilation will be maintained when disease affects one or more afferent pathways or when the perception of some sensory cue is blunted by anesthesia, sleep, or neural injury. Conflicting demands, signals, or both, from different receptors may be responsible for dyspnea, a common symptom in respiratory disease, or for hiccups, cough, and postural events that also utilize respiratory muscles. The system is also driven by feed-forward mechanisms. One example is the anticipation of ventilation that may precede exercise even in the absence of CO2 sensitivity [52]. Feed-forward mechanisms play a critical role in smoothly integrating breathing with swallowing, speaking, singing, and defecating. Pontine and medullary respiratory groups initiate ventilatory behavior [53]; these areas regulate the firing and pattern of discharge in the motor
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Cortical State O2 Consumption Hypothalamic Regulation
CO2 Production
Pontomedullary Outflow Neural Transmission
Weight Medulla (VMS) Carotid Body
Frequency
Muscle Controlled Variables: O2 content, pH
Minute Ventilation
Tidal Volume
Measured Variables
Figure 8.2 This diagram identifies some of the anatomic and functional elements in the creation of ventilatory behavior. It is very simplistic. The controller elements are the cortex, pons and medulla; the controlled elements are the neuromuscular and anatomic elements of the upper airway and chest wall; the controlled variables are identified as arterial levels of CO2 and O2. Variables that are measured in studies of unanesthetized and unrestrained animals are noted. Weight is a loading factor to the controlled elements, but also is a symbol of metabolic influences on ventilatory behavior through hypothalamic function.
neurons of chest wall and upper airway muscles in a sequential and nonrandom fashion. Discharge patterns in some pontine cells appear to be dependent on afferent vagal feedback but become more clearly phasic after vagotomy. Thus, in the adult mammal neither the medulla nor the pons alone generates the respiratory pattern. In addition, there is an interaction between states of alertness and the activity of higher brain centers and the brain stem bulbopontine respiratory neurons that determines respiratory rhythm and depth. A detailed discussion of the theory and the experimental data on the generation of respiratory rate and depth [53] is beyond the scope of the current review; however, the notion that there are several current models is important if one is to think of linkage between genes and ventilatory behavior. Electrical (neural) and mechanical outputs of the central inspiratory activity involve the anatomic structures of the chest wall (to inflate the lungs) and the upper airway (to stabilize the air channel to the lungs) [7]. In the diaphragm there is a progressive rate of rise in inspiratory activity
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that allows the musculature to overcome the progressive increase in the elastic recoil of the lung during inflation. The rate of rise of central inspiratory activity, and hence the rate of lung inflation, is controlled by mechanisms different from those that terminate inspiration. Hypoxia and hypercapnia increase the steepness of the ramp of inspiratory activity and hence increase the rate of inspiratory airflow and tidal volume, but they have little effect on the duration of inspiration and, therefore, on the breathing frequency. This means that ventilatory responses to hypercapnia and hypoxia will depend on the sensitivity of both chemoreceptors and stretch receptors. Chemoreceptor sensitivity, since it influences the rate of increase in central inspiratory activity, is related more closely to the level of average airflow during inspiration than to minute ventilation. The change in the tidal volume/inspiratory time ratio, rather than the change in ventilation itself, more closely reflects chemical drive. On the other hand, the change in inspiratory time as a fraction of total breath duration may relate to afferent feedback such as that from the activity of stretch receptors. Additional factors alter the effect of respiratory central drive on tidal volume and frequency. With diseases of the lungs and chest wall, with obstructive apnea during sleep, or during certain stages of sleep when nonrespiratory drive is altered in muscles of the chest wall, there can be a mismatch between neural activity and mechanical result [7]. Hence, features of the controlled elements of the chest wall and upper airway can obscure central drive, especially if the intervention substantially alters the controlled system. The choice of using a single or double chamber for the barometric methods is in part determined by whether one suspects lung mechanics to substantially change during testing [54], since tidal volume and inspiratory flow are not always a precise copy of central drive. Ideally, one would pick a collection of phenotype traits that are sensitive to single features of ventilatory behavior; however, it is not feasible to examine all parts of respiratory control at once. The need to study large numbers of small animals and at times permit these test subjects to breed has necessitated the use of non-invasive measures of ventilation, namely tidal volume and frequency, using the barometric method (Figure 8.3) [55]. There is anguish over this approach [54]. Most agree that frequency is a very precise value, but tidal volume is semi-quantitative [60]. The latter is estimated from temperature changes which in turn can be affected by the mammal’s temperature and the temperature and humidity of the chamber [55,61]. A major strength of this approach is the ability to measure an animal more than once [62]. Variability is induced by day-to-day alterations in such environmental factors as light, smell, noise, etc., but can be reduced by careful attention to the animal protocol [54]; time of day also influences trait values [63,64]. These variables enter into estimates
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Zucker fatty F2 F2BN 1 2 3 4 5 6 7* 8 9
F1
Key: fa−
fa+
950 bp 700 bp
3 = fa/fa 4 = fa/fa 6 = fa/− 7*=-/-
F2 1/4
1/2
1/4
Fatty
Figure 8.3 An intercross strategy is shown for a cross between the Zucker and the Brown Norway rat. The allele illustrates the transmission of the mutant fatty (noted as faþ) gene. One estimates the strength of inheritance by the difference in the variance in trait values among the F1 generation (which is genetically identical and whose variance is environmental) and the variance in the F2 generation (where the genes are mixed and the environmental component is assumed to be the same as with the F1 animals). Such estimates can be made for animals in the F2 generation that carry the faþ and/or fa null (fa) allele.
of the environmental component of the protocol, and if too large or uncontrolled may obscure a genetic effect. Hence, studies of unanesthetized animals require larger numbers or statistical adjustments that reduce such influences. Reduced preparations or anesthetized models require more considerable attention to reproducibility of trait values. In anesthetized mice there is good correspondence between estimates of pulmonary mechanical problems identified by the barometric method when compared with more direct measures of mechanics like force oscillatory techniques [65]. Therefore, one can conceive of approaches that compare animal strains in regard to dose– response relationships in chemical drive, or the anatomic or physiologic characterization of brain regions and/or the individual influences of resistance and compliance of upper airway and chest wall structures on total respiratory mechanics. However, the exclusive use of reduced preparations and more technologically advanced approaches is appropriate for times in which the genes or proteins are better characterized. The use of a reduced preparation is most convincing in regard to evolutionary fitness when used in tandem with other methods to disclose mechanisms for genetic effects [66]. Another source of worry is the reproducibility of results across laboratories. There is some empiric evidence for this general concern, although not directly related to ventilatory behavior. After standardizing strain source and testing systems, there were systematic differences in
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behavior across laboratories in a battery of six tests for neurobehavior. For some tests, the magnitude of difference ascribed to strain depended upon the specific testing lab [67]. Thus, care must be taken in characterizing the behavior in both common strains and mutants, as results could be idiosyncratic to a particular laboratory. At present there is no such comparable study for ventilatory behavior. There is some data to suggest that absolute values for such traits as frequency and tidal volume vary between laboratories that use slightly different methods of testing. Despite such differences, the ranking of animals to each other, in particular the pattern of breathing for the C57BL/6J, is similar between laboratories [68–71]. Animal models of human problems like sleep apnea exist in larger animals (dogs, elephant seals, and pigs) but the rodent models (mice and rats) are amenable to genetic dissection. Studies in these models can address environmental exposures, consequences, and intermediate phenotypes found in humans, which may relate to OSA expression or progression [72–74]. Utilizing inbred strains reduces genetic variability and permits more control over age effects and environmental factors that confound human studies. Simultaneous use of proteomics and expression arrays in such models may efficiently screen many biochemical processes that could contribute to trait variance, and can be useful in locating candidate genes within the regions-of-interest identified through the study of in-bred lines. This approach is used to identify genomic elements operating in strain differences in sleep in the mouse [75]. Complementary and parallel studies in animals and humans can be more efficient in understanding the biology of sleep-disordered breathing and sleep hypoxemia when compared with a sequential approach. Compared with the mouse, the rat’s larger size and betterdefined brain nuclei are amenable to follow-up studies of cardiovascular and respiratory control in the central nervous system. Respiratory mechanics of the upper and lower airways are in relative proportion more similar between the rat and the human than between the human and the mouse [2]. If one considers a need for arterial blood gas measurements as a phenotype trait, one would have to consider not only the technical accuracy and reproducibility of the measurement but also the variability in the state of the animal. Both effects limit the power of intermittent sampling of arterial blood gases to detect differences in gas exchange or acid–base between strains or in their progeny. Technology for on-time sampling of gas exchange, particularly if such methods were non-destructive, might be useful. Finally, the laboratory must be directed toward a goal of having reproducible testing conditions over the life of the rodent study or cross [76]. Changes in testing methods or personnel, and in animal quarters, may all obscure detection of small (53%) but important gene effects. In short,
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the more uniform the testing conditions, the less influence there is for environmental variance and the more likely one will detect genetic effects and interactions. Identification of gene loci that link to intermediate traits has given insight into risk factors for cardiovascular and metabolic disease [77,78]. The start of this field in regard to respiratory control was the empirical observation that ventilatory phenotype was a factor in the variations among mouse strains in the model of environmental ozone exposure [73]. What is needed is the examination of other models, including those gene knock-outs for cardiovascular or neurologic functions that might operate in the risk for or recognition of sleep apnea [79]. A survey of respiratory frequency at rest and with chemoresponsiveness could provide insight into markers for risk of unexpected death in infancy or for screening adults with hypoventilation complications with illnesses like sleep apnea, COPD, and asthma [78,80,81], or lethal human illnesses such as idiopathic or congenital hypoventilation syndromes [32,82]. In summary, the key to any approach to the genetics of respiratory control is an intimate knowledge of the trait value and the animal. Initial results are likely to be confounded by sources of variance and by major confounding variables. The assignment of gene action linked to ventilatory behavior to a particular candidate physiologic process should be a statistical or empirical rather than an intuitive process. Bias introduced at every level of collection of the phenotype data may enter into analysis and produce or obscure linkage. The physiologist will need to prove suspected biologic linkage in any number of ways [6]. Nevertheless, in this postgenome era there is enormous opportunity to measure ventilatory traits in any number of mammalian models and to begin to describe disorders of respiratory control in terms of molecules relevant to whole system needs, like ventilation, metabolism, and responses to sleep and chemosensory stress. IV.
Evidence for the Inheritance of Ventilatory Traits in Rodents
In 1984, Ou et al. [83] reported differences in the ventilatory behavior between two colonies of Sprague-Dawley rats in response to chronic exposure to hypoxia. Despite these being out-bred strains (no conscious in-breeding by brother–sister pairing for 420 generations), consistent differences in the ventilatory response to hypoxia before the exposure remained between colonies over the course of several years, and correlated to differences in the erythropoietic and pulmonary vascular pressor responses to chronic hypobaric hypoxia [84]. By the vendors maintaining such closed populations, there had occurred genetic drift between the
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populations. The problem with the study of these Sprague-Dawley strains is that the similarities between the genomes are so great that finding the responsible gene is extremely difficult. In 1992, Connelly et al. [85] reported (almost as an afterthought) the observation that the effect of N-methyl-D -aspartate (NMDA) receptor-channel activation in the production of respiratory patterns differed by strain in an anesthetized rat model. The predominant respiratory response to systemic MK-801 administration, a pharmacologic blocker of NMDA activation, was an increase in inspiratory duration and a decrease in amplitude of diaphragm electromyogram and phrenic nerve discharge. Effects on inspiratory timing and amplitude were most pronounced when the rats were vagotomized. Respiratory timing changes in response to systemic MK-801 administration differed between the Wistar and SpragueDawley strains. Breathing patterns resembling apneusis, i.e., irregular inspiratory durations prolonged two- to thirty-fold, occurred in 60% of the vagotomized, spontaneously breathing Sprague-Dawley rats and in none of the Wistar rats. The authors concluded that the breathing pattern in Sprague-Dawley rats is more sensitive to interference with NMDA-mediated mechanisms. This study is important as it illustrates that strain differences may occur in ventilatory behavior even in the anesthetized preparation, so that models of the neurochemistry of breathing based upon findings from one strain need to be tested for face validity in another strain. In 1995, Tankersley et al. [73] reported results from the more traditional approach of surveying many in-bred strains, in this case mice, in order to detect if any strains showed significant divergence in trait values. There were significant inter-strain differences in respiratory frequency, hypoxic responsiveness, and hypercapnic responsiveness between several strains [70]. Other reports from this group indicate success in an intercross strategy between C57BL/6J (B6) and C3H/HeJ (C3) to uncover gene regions involved in ventilatory behavior. This group has subsequently reported, in a similar intercross, that a strain distribution pattern for respiratory rate when exposed to hypoxia is consistent with the hypothesis that two genes regulate parental strain differences [86,87]. Cosegregation analyses suggest that the genetic control of frequency during hypoxia differs from the genes that control differential baseline frequency [88]. Although the genetic control of tidal volume appears more complex, differences in the minute ventilation during hypoxia is determined by tidal volume responses. In this instance, the major gene regions that may relate to lung mechanics appear to be distinct from those correlated to inspiratory drive [89]. Therefore, this study suggests that the phenotypic variation in hypoxic ventilatory response between the two parental strains, especially related to frequency
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during hypoxia, is possibly regulated by only a few major genetic determinants. In 1997, Strohl et al. [90] reported measurements of ventilation and metabolism in four strains of rats, chosen for a wide variation in traits for body weight and/or blood pressure regulation. The conclusion is that strain, more than the effect of body mass or sex, had a major influence on metabolism, the pattern and level of ventilation during air breathing, and ventilation during loading or unloading of chemoreceptor input in the unanesthetized rat. A more recent study found similar results in four strains of rat, including a confirmation of the low CO2 response in the Brown Norway [91]. These strain differences indicate that the ventilatory response to chemosensory input appears to be genetically regulated. There are indications that inheritance operates in some of the nonsteady-state responses. The Dejours phenomenon is very clearly demonstrated under non-steady-state conditions involving rapid transitions from room air to hypoxia and then to hyperoxia. This transition is quite complex when compared with steady-state responses to hypoxia and/or hypercapnia and, at least intuitively, is thought to be due to an unnatural environmental exposure. To test this null hypothesis, we exposed two rat strains known to differ in ventilatory behavior to oxygen after exposure to hypoxia. We examined the ventilatory behavior (frequency, tidal volume and minute ventilation) in Sprague-Dawley and Brown Norway animals. The phenomenon of ventilatory decline in response to 100% oxygen after 5 min of exposure to hypoxic gas (10% oxygen, 90% N2) was not common in the Sprague-Dawley strain, but was quite evident in the Brown Norway strain (Figure 8.4). This study rejected the null hypothesis and suggests that genetic factors do influence the ventilatory response to hyperoxia following acute hypoxic exposure [92]. Such non-steady-state responses could be important intermediate traits relevant to recurrent cycles of hypoxia-reoxygenation present in sleep-disordered breathing. The presence of post-hypoxic ventilatory decline is found in those patients with heart failure and Cheyne–Stokes respiration, rather than those without Cheyne–Stokes respiration [93]. The above studies in common strains of mice and rats offer the most compelling argument for ventilatory behavior being affected by genetic drift and natural selection and offer the foundation for physiological genomic studies aimed at elucidating the genetics of these ventilatory control mechanisms. As will be discussed below, the effects seen in these models can be quite modest, and the assignment of cause-and-effect relationships can be complex given that adjustments in one gene may influence another gene or gene product and one gene can have an influence on more than one trait or part of the system for respiratory control.
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SUCTION Monitor Temp., humidity , %O2 , %CO2
Reference chamber
GAS MIXTURE
Ventilatory behavior (freq, TV, Ve)
PRESSURE TRANSDUCER
Figure 8.4 This diagram illustrates the elements of a barometric method for measuring ventilatory behavior in rodents (Data from Ref. 90). Not shown in detail is the use of a sealed reference chamber, identical in size as the animal chamber.
V.
Estimates of the Strength of Inheritance
The techniques used to examine experimental crosses are now more powerful and flexible [6]. Current approaches can accommodate environmental influences, make minimal assumptions about the mode of inheritance of the trait, and can accommodate effects of factors such as age, growth, and cohort effects, unknown ascertainment, heterogeneity, and heterozygosity in parental lines. Observational data should include real or potential temporal changes in litter size, food vendor and brand of rat chow. These quantitative and qualitative variables are handled as co-variate shared environmental variables. Animal husbandry reduces heterozygosity and the intercross strategy will provide a reliable means for estimating environmental effects and permit heritability estimates for a variety of important ventilatory phenotypes [94]. For the assessment of the gross effects of genes and environmental factors on phenotypic values, variance components models are used [94]. The total phenotypic variance of each trait is partitioned into a genetic
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Sprague Dawley
insp
Air
10% O2
100% O2
5thmin
1stmin
Brown Norway
time
1999
Figure 8.5 The barometric tracings from two animals (one Sprague-Dawley and one Brown Norway) are compared with regard to the pattern of breathing at rest, in the fifth minute of hypoxia, and in the first minute or so after re-oxygenation from hypoxia (adapted from Ref. 92).
component and an environmental component. One then estimates the strength of inheritance by the difference in the variance in trait values among the first generation animals (which is genetically identical and whose variance is environmental) and the variance in the second generation (where the genes are mixed and the environmental component is assumed to be the same as with the first generation animals). Experience with cross-breeding of the Sprague-Dawley and Brown Norway and of the Brown Norway and the Zucker strains has permitted us to make such calculations. A schematic diagram of the strategy for this cross is presented in Figure 8.5. A summary of the estimates for some traits for ventilatory behavior is presented in Table 8.2. In this instance, estimates are provided from analyses of the variance [95] in first- and second-generation animals of a cross between a Zucker fat male and Brown Norway females. From this data we suggest that some but not all trait values related to ventilation and metabolism show higher (440%) inheritance. The strength of inheritance for some ventilatory traits is similar to that described for airway responsiveness in mice [61].
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Table 8.2 Heritability Estimates from a Cross between the Zucker and Brown Norway Strains
Trait Weight Lee index (mass) Carbon dioxide production Frequency at rest Minute ventilation Frequency in hypercapnia Ventilation in hypercapnia Frequency in hypoxia Minute ventilation in hypoxia
Total heritability (h2)
Variance attributed to the fa allele
Variance attributed to the rest of the Genome
0.78 0.72 0.55 0.59 0.11 0.28 0.38 0.60 0.54
47% 38% 10% 9% 6% 17% 0% 11% 0%
53% 62% 90% 91% 94% 83% 100% 89% 100%
The Zucker rat strain carries a mutant allele, the fa, that alters the leptin receptor; two such mutant alleles results in an obese phenotype due to leptin resistance (Data from Ref. 129).
VI.
Studies of Gene Effects in Rodent Models
If one were to identify the best evidence for genetic effects, it would be in the literature on genetically engineered animals and studies of drug-by-strain interactions. These studies indicate that genetic background directly influences ventilatory behavior (respiratory frequency, tidal volume, and/ or minute ventilation) either at rest or with hypercapnic challenges. There is evidence demonstrating the action of genes that arises out of studies of nitric oxide synthase (NOS). There are three isoforms of NOS (NOS-1, NOS-2, and NOS-3) arising from three different regions of the genome, and these enzymes are involved in the generation of nitric oxide (NO). The concept of genetic transmission of post-hypoxia ventilatory behavior is directly supported by reports of nearly absent respiratory depression in response to brief hyperoxia in nitric oxide synthase (NOS)-3 mutant mice [96]. In addition, an altered breathing pattern was seen in NOS-1 mice where function was eliminated by knock-out [97,98]. Other studies of knock-out (KO) mouse models report loss of or reduction in hypoxic response attributed to endothelin-converting enzyme-1 (ECE-1) [99], to endothelin-1 [100], to dopamine [101], to the neutral endopepidase (NEP) [102], and to HIF [103]. On the other hand, animals knocked-out for other supposedly critical factors for hypoxic or hypercapnic responses show no effect on ventilation; this is the case for endothelin-3 [104]. Some genes may have functions in developmental programs. The reports on the NK-1 KO suggest that NK1 receptors are important in the response to acute hypoxia in the adult mouse; however, NK1 receptors
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are not obligatory for the prenatal development of the respiratory network, for the production of the rhythm, or for the regulation of breathing by short-lasting hypoxia in neonates [105]. When comparisons are made between monoamine oxidase A-deficient transgenic (Tg8) mice with control (C3H), the implication is that an elevated serotonin during the perinatal period alters respiratory network maturation and produces a permanent respiratory dysfunction, whereas a high serotonin level present in adults depresses chemosensation; the authors concluded that the metabolism of serotonin plays a crucial role in the maturation of the respiratory network and in ventilatory response to hypoxia [106]. The NADPH system is of interest for its presumed role in oxygen sensing at a mitochondrial level. There is a role for the subunit of NADPH, gp91, in cellular sensing of oxygen in airway neuroepithelial cells [107], but not in adrenal cells [108]. Yet in KO models there is little effect on pulmonary vascular [109] or carotid body [110] cellular responses to hypoxia, or in ventilatory behavior [111]. In contrast to the interest in hypoxia, there is less work in models on chemoresponsiveness to CO2. Development of carbon dioxide responses is impaired in KO models for the mammalian achaete-scute homologous gene (Mash-1, now called Ascl 1) [112]. Adult monoamine oxidase A-deficient transgenic mice, when compared with control (C3H) mice, show blunted CO2 responses which are restored when serotonin levels are pharmacologically suppressed [106]. One interesting study that utilized CO2 loading was able to disprove the null hypothesis that exposure to high levels of carbon dioxide would not result in differences in early-gene (c-fos) expression among strains [113]. This approach probably raises more questions than it answers, but in short it does document the possibility for intra-species variations in the widespread brain networks that could act not only on ventilatory behavior but on acid–base balance as well [114]. A more complex model that is still based on single mutant genes includes the ob/ob mutant mouse [86,115,116]. This model is based upon the C57BL/6J strain and is a good example of a modifier gene effect. The basic structure of resting breathing is unaffected but the function of carbon dioxide responses is modified by the leptin system, even in the absence of obesity. Thus, a candidate gene/protein approach has shown proof-of-principle for providing insight into functions of individual genes relevant to ventilatory behavior and human conditions like obesity and sleep-disordered breathing. Knock-out models, while of significant interest, are often not relevant to understanding risk factors since evolutionary fitness requires actions of both the real and latent variance in several genes and gene products, respiratory system capacity, and learning. Understanding the role of genes
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in normal and pathologic control of ventilation will require approaches that will accommodate the probable collective effects of multiple loci. Another way to illuminate genetic influences is to examine drug effects in different strains of rodent. Since studies in NOS KO mice have shown that the nitric oxide system can influence post-hypoxic ventilatory behavior [66,96,117], the question arises whether nitric oxide synthase (NOS) blockade would similarly affect ventilatory behavior in two rat strains. Post-hypoxic decline in frequency and minute ventilation (the Dejours phenomena) is present in some but not all studies of Sprague-Dawley rats, a finding that might relate to colony differences [92]. In Sprague-Dawley animals without this response, global NOS blockage produced the Dejours phenomenon, while NOS blockade resulted in a greater increase in resting ventilation in the Sprague-Dawley than in the Brown Norway strain, and uncovered the Dejours phenomenon in the Sprague-Dawley strain [118]. Based on this observation we proceeded to administer a selective neuronal NOS (NOS-1) blocking agent (7-nitroindazole or 7-NI, 60 mg/kg) and found the same effect as the non-specific NOS inhibitor but without altering metabolism (unpublished findings). These findings are consistent with the effect of NOS inhibition in the control mice studied in relation to their NOS-1 KO cousins [98]. These approaches are intended to determine if a specific system, in this case the nitric oxide system, operates similarly in all rodents. As the results indicate, the various strains of rats and mice do not utilize the NO pathway in the same way with regard to the regulation of ventilatory behavior. The strength of the knock-out approach or drug-by-strain approach is the comfortable connection between a known gene, peptide, or protein and the ability to make a rational connection to ventilatory behavior. Falsepositive findings may occur due to selection bias and/or over-interpretation of the significance of results from knock-out models and/or reduced preparations, especially if one uses only this information to screen human populations. Multiple approaches are preferable, and there are a number of incremental steps that must be taken before one can assign a specific role to a gene. The pioneering work by Tankersley and his group has identified some gene regions that are statistically linked to differences in ventilatory behavior between C57BL/6J and C3H/HeJ mice [74,88,119,120]. Quantitative statistics of animals derived from these strains suggest that the genetic control of hypoxic ventilatory responses exhibits a relatively simple Mendelian inheritance in terms of respiratory timing characteristics. Furthermore, differences in inspiratory time at baseline is linked to a putative genomic region on mouse chromosome 3 [88]; however, a likely candidate gene related to neuronal receptors does not explain this association [121]. Of interest, this region does not correlate with the variations in lung mechanics between these two strains [89]. This literature
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encourages the use of functional genomics in an effort to link ventilatory behavior to molecular structure [120]. VII.
A Physiogenetic Map of Ventilatory Behavior
A literature on the genetics of ventilatory behavior is emerging with animal models and traits that appear relevant to human physiology and pathophysiology. One theme is that a limited number of genes can have a measurable impact on ventilatory behavior. The overview of the general idea of how genes act to produce the ventilatory phenotype is illustrated in Figure 8.6. From this perspective it is remarkable that studies of ventilatory behavior have identified any genetic effects (Table 8.3). It is likely the genes that ultimately influence ventilation at rest may also affect ventilation when breathing is stimulated or may correlate with regulatory systems for the cardiovascular system, and perhaps even metabolism. This makes sense when one considers how well integrated, from a physiological standpoint, the systems and mechanisms that regulate respiration are. It will, therefore, be important to consider the presence of gene interactions in the data analysis. Such interactions are revealed not only as preliminary data for the mapping of gene loci, but also as a way of gaining insight into the interrelationships and nature of the genetic mechanisms influencing ventilatory responsiveness [122]. There are also patterns of interactions among the physiologic traits we use for describing ventilatory behavior, and associations that differ among
Ventilation during sleep and wakefulness
f
TV
SYSTEMS LEVEL
………. ………
CELL LEVEL
Genes or Gene Regions
Figure 8.6 The pathways between genes and the ventilatory traits of respiratory frequency ( f ) and depth (tidal volume or TV) are complex and involve several levels of cellular and system integration.
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Table 8.3 Candidate Genes or Regions Identified (February 2003) Gene
Human region
ACE POMC APOE Unknown NOS-1 neuronal NOS-3 (eNOS) Unknownd
17, q23 2, p23 19, q13.2 8, 137cMc 12, q24.2 7, q36 4, 3, and 13
Unknown Endothelin-1 (Edn-1) ECEC-1 HIF-1a Dopamine (1a) receptor Monoamine oxidase A Ascl 1
6 and 15 6, p24.1 6, q36–37 14, q21–24 11, q11 X, p11.4–11.3 12, q22–23
Mouse region 11 (65 cM) 14 (4 cM) 7 (4 cM) Unknown 5 (65 cM) 5 (5 cM) D3mit7 (approximately 26.4 cM) (site of ob mutation in B6 strain) D9mit207 13, 26 cM Unknown 12, 31 cM 9 (drd2) X, 5.2 cM 10 (ascl1)
Referencea Speciesb [40,41] [128] [28] [128] [98] [96] [88]
Human Human Human Human Mouse Mouse Mouse
[87] [100] [99] [103] [101] [106] [130]
Mouse Mouse Mouse Mouse Mouse Mouse Mouse
a
Primary observation(s). Species in which the observation was first made. c No gene mapped to this region as yet. d Contributes to the strain differences to susceptibility to diet (fat) induced obesity. Source for homology information: http://www.informatics.jax.org/. b
strains. Figures 8.7a and b result from our work in the Sprague-Dawley and Brown Norway rat strains and a collaboration with Physgen, the Program in Genomic Application, based at the Medical College of Wisconsin, and headed by Dr. Howard Jacob. Even in a grey scale, the eye can detect pattern differences clustering among traits in the Sprague-Dawley (SD) strain. Also the eye can detect differences in these patterns between SpragueDawley and Brown Norway animals (n ¼ 55 in each group). This approach results in the creation of a systems map for ventilatory function [122]. This map can be used alone or in combination with whole genome mapping to disclose interactions among ventilatory traits. Some effects that may be disclosed are pleiotropy, epistasis, and sexual dimorphism [94]. Pleiotropy, one gene or the mutation of a gene resulting in the production of apparently unrelated (multiple) effects at a phenotypic level, has been observed by us in an association between values for resting breathing and hypoxic ventilatory response in humans [123]. Epistasis is an interaction of two or more loci on any given trait value; more technically, it is a form of gene interaction whereby one gene interferes with the phenotype
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Bivariate Coefficients of 24 Traits
Metabolism Resting Breathing 5thmin. Hypoxia 5thmin. Hypercapnia 10% O2 Responsiveness Hypercapnic Resp. Isocap. Hypox Resp. Re-oxygenation (Dejours)
CORRELATIONS DIFFER BETWEEN STRAIN
Figure 8.7 (a) In this example we performed Pearson coefficient correlations among the 24 traits comprising eight groups of systems actions in a group of Sprague-Dawley animals (n ¼ 55). The correlation coefficient is presented as a set of colors (shown here in grey scale) from þ1 right hand of the scale to 1. The original graphs are color codes; however the structure is disclosed by grey-scale. (b) Shown for comparison are the correlations for the Sprague-Dawley (Figure 8.7a) and the Brown Norway (n ¼ 55). These figures were constructed as examples of physiogenomic approaches in collaboration with Peter Tonellato and the Program for Genomic Application, the Medical College of Wisconsin (see Ref. 108 for other examples).
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expression of another non-allelic gene [94]. It is a mechanism by which two loci can have more (or less) than an additive effect. Sexual dimorphism occurs in many physiologic variables and can occur at both chromosomal or hormone environment level. Imprinting refers to a differential effect, at either a chromosomal or allelic level, depending upon whether the genetic material came from a male or female parent. A clinical example is transmission of either Prader-Willi or Angelman syndrome, human conditions associated with respiratory disturbances during sleep [124–127]. Only gene mapping proves this effect, but some inference about its action can be gained by pedigree analysis. VIII.
Overview and Future Directions
Results of these studies of inheritance of ventilation, tidal volume, and frequency may permit the design of appropriate experiments including rational decisions to examine the physiologic systems for the loci identified by gene mapping. If results indicate a substantial effect in respiratory mechanics, attention could be directed to specific biochemical, morphometric, and mechanical features. More conclusive investigation would include the more complicated measures of such things as the mechanical system (controlled element), arterial blood gases (controlled as well as controlling element), and upper airway mechanics (controlled element). Identification of trait influencing loci will be relevant to understanding diseases characterized by either hypo- or hyper-ventilation and/or by alterations in respiratory patterning [101]. We would have greater opportunity to understand the heterogeneity in responses to environmental influences and, potentially, to develop a set of markers that helps address breathing patterns in adults with hypoventilation or with illnesses like congestive heart failure, COPD, and asthma. However, it is unrealistic to propose that genetic factors explain all of the variability in the expression of ventilatory behavior or its pathogenic effects on cardiopulmonary disorders of sleep. Rather, the questions currently being addressed relate to the relative strength of the genetic components and the identification of the genes, proteins, and/or systems that produce ventilatory dysrhythmia and the fundamental nature of the periodic hypoxemia found in sleep-disordered breathing. A reasonable expectation related to therapeutic utility would be the identification of disease-modifying genes, including those that amplify a given stimulus, sensitize (desensitize) an effector pathway, or promote a stimulus–response mismatch. Having such markers will permit the design of subsequent studies to identify individuals at greater or lesser risk. Such stratification would complement current clinical practice and begin efforts to identify those at
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risk, predict morbidity, or otherwise inform in regard to appropriate surveillance or clinical management in cardiopulmonary illnesses in which sleep-disordered breathing occurs. Acknowledgments The author would like to thank the Department of Veterans Affairs and the National Heart, Lung and Blood Institute for support of this line of research. The author was a visiting Scientist to the Program for Genomic Application, the Medical College of Wisconsin (Howard Jacob, Ph.D., Director) as part of a Sleep Academic Award program from the National Institutes of Health. References 1. 2.
3. 4. 5. 6. 7. 8. 9.
10.
11. 12. 13.
Agostoni, E., Thimm, F. and Fenn, W.O., Comparative features of mechanics of breathing, J. Appl. Physiol. 14, 679–683, 1959. Crossfill, M.L. and Widdicombe, J., Physical characteristics of the chest and lungs and the work of breathing in different mammalian species, J. Physiol. 158, 1–14, 1961. Bennett, F.M. and Tenney, S.M., Comparative mechanics of mammalian respiratory system, Respir. Physiol. 49, 131–140, 1982. Cherniack, N.S. and Altose, M.D., Mechanisms of dyspnea, Clin. Chest. Med. 8, 207–214, 1987. Strohl, K.P. and Thomas, A.J., Breathlessness, anxiety, and respiratory physiology, Psychosom. Med. 60, 680–681, 1998. American Thoracic Society, Finding Genetic Mechanisms in Syndromes of Sleep Disordered Breathing, www.thoracic.org, 1999. Cherniack, N.S. and Longobardo, G.S., The chemical control of respiration, Ann. Biomed. Eng. 9, 395–407, 1981. Strohl, K.P., Cherniack, N.S. and Gothe, B., Physiologic basis of therapy for sleep apnea. Am. Rev. Respir. Dis. 134, 791–802, 1986. Shea, S.A., Horner, R.L., Benchetrit, G. and Guz, A., The persistence of a respiratory ‘‘personality’’ into stage IV sleep in man, Respir. Physiol. 80, 33–44, 1990. Shea, S.A., Dinh, T.P., Hamilton, R.D., Guz, A. and Benchetrit, G., Breathing patterns of monozygous twins during behavioural tasks, Acta Genet. Med. Gemellol. (Roma) 42, 171–184, 1993. Shea, S.A. and Guz, A., Personnalite´ ventilatoire — an overview, Respir. Physiol. 87, 275–291, 1992. Beral, V. and Read, D.J.C., Insensitivity of respiratory center to carbon dioxide in the Enga people of New Guinea, Lancet 2, 1290–1299, 1971. Saunders, N.A., Leeder, S.R. and Rebuck, A.S., Ventilatory response to carbon dioxide in young athletes: a family study, Am. Rev. Respir. Dis. 113, 497–502, 1976.
284 14.
15. 16. 17.
18.
19.
20.
21.
22. 23.
24.
25.
26.
27.
28.
29.
Strohl Scoggin, C.H., Doekel, R.D., Kryger, M.H., Zwillich, C.W. and Weil, J.V., Familial aspects of decreased hypoxic drive in endurance athletes, J. Appl. Physiol. 44, 464–468, 1978. Collins, D.D., Scoggin, C.H., Zwillich, C.W. and Weil, J.V., Hereditary aspects of decreased hypoxic response, J. Clin. Invest. 70, 105–110, 1978. Kawakami, Y., Yoshikawa, T., Shida, A., Asanuma, Y. and Murao, M., Control of breathing in young twins, J. Appl. Physiol. 52, 537–542, 1982. Kawakami, Y., Yamamoto, H., Yoshikawa, T. and Shida, A., Age-related variation of respiratory chemosensitivity in monozygotic twins, Am. Rev. Respir. Dis. 132, 89–92, 1985. Kawakami, Y., Shida, A., Yoshikawa, T. and Yamamoto, H., Genetic and environmental influence on inspiratory resistive load detection, Respiration 45, 100–110, 1984. Kawakami, Y., Yamamoto, H., Yoshikawa, T. and Shida, A., Respiratory chemosensitivity in smokers. Studies on monozygotic twins, Am. Rev. Respir. Dis. 126, 986–990, 1982. Kawakami, Y., Yamamoto, H., Yoshikawa, T. and Shida, A., Chemical and behavioral control of breathing in twins, Am. Rev. Respir. Dis. 129, 703–707, 1984. Kobayashi, S., Nishimura, M., Yamamoto, M., Akiyama, Y., Kishi, F. and Kawakami, Y., Dyspnea sensation and chemical control of breathing in adult twins, Am. Rev. Respir. Dis. 147, 1192–1198, 1993. Kawakami, Y., Shida, A., Yamamoto, H. and Yoshikawa, T., Pattern of genetic influence on pulmonary function, Chest 87, 507–511, 1985. Kawakami, Y., Irie, T., Shida, A. and Yoshikawa, T., Familial factors affecting arterial blood gas values and respiratory chemosensitivity in chronic obstructive pulmonary disease, Am. Rev. Respir. Dis. 125, 420–425, 1982. Moore, G.C., Zwillich, C.W., Battaglia, J.D., Cotton, E.K. and Weil, J.V., Respiratory failure associated with familial depression of ventilatory response to hypoxia and hypercapnia, N. Engl. J. Med. 295, 861– 865, 1976. Kafer, E.R. and Leigh, J., Recurrent respiratory failure associated with the absence of ventilatory response to hypercapnia and hypoxemia, Am. Rev. Respir. Dis. 106, 100–108, 1972. Redline, S., Leitner, J., Arnold, J., Tishler, P. and Altose, M., Ventilatory control abnormalities in familial sleep apnea, Am. J. Respir. Crit. Care Med. 156, 155–160, 1997. Redline, S. and Tishler, P., Familial influences on sleep apnea, in Lung Biology in Health and Disease: Sleep-Related Breathing Disorder, Lenfant, C. and Saunders, N., eds., New York, Marcel-Dekker, pp. 363–378, 1994. Kadotani, H., Kadotani, T., Young, T., Peppard, P.E., Finn, L., Colrain, I.M., Murphy, G.M., Jr. and Mignot, E., Association between apolipoprotein E epsilon4 and sleep-disordered breathing in adults, JAMA 285, 2888–2890, 2001. Saarelainen, S., Lehtimaki, T., Kallonen, E., Laasonen, K., Poussa, T. and Nieminen, M.M., No relation between apolipoprotein E alleles and obstructive sleep apnea, Clin. Genet. 53, 147–148, 1998.
Inheritance and Ventilatory Behavior in Animal Models 30.
31.
32.
33.
34.
35.
36. 37.
38.
39.
40.
41.
42.
43. 44.
285
Foley, D.J., Masaki, K., White, L. and Redline, S., Relationship between apolipoprotein E epsilon4 and sleep-disordered breathing at different ages, JAMA 286, 1447–1448, 2001. Buxbaum, S.G., Elston, R.C., Tishler, P.V. and Redline, S., Genetics of the apnea hypopnea index in Caucasians and African Americans. I. Segregation analysis. Genet. Epidemiol. 22, 243–253, 2002. Tishler, P.V., Redline, S., Ferrette, V., Hans, M.G. and Altose, M.D., The association of sudden unexpected infant death with obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 153, 1857–1863, 1996. Palmer, L.J., Buxbaum, S.G., Larkin, E., Patel, S.R., Elston, R.C., Tishler, P.V. and Redline, S., A whole-genome scan for obstructive sleep apnea and obesity, Am. J. Hum. Genet. 72, 340–350, 2003. Wilcox, I., McNamara, S.G., Collins, F.L., Grunstein, R.R. and Sullivan, C.E., ‘‘Syndrome Z’’: the interaction of sleep apnoea, vascular risk factors and heart disease, Thorax 53, S25–S28, 1998. Marks, D.L. and Cone, R.D., Central melanocortins and the regulation of weight during acute and chronic disease, Recent Prog. Horm. Res. 56, 359–375, 2001. Ukkola, O. and Bouchard, C., Clustering of metabolic abnormalities in obese individuals: the role of genetic factors, Ann. Med. 33, 79–90, 2001. Watson, P.M., Commins, S.P., Beiler, R.J., Hatcher, H.C. and Gettys, T.W., Differential regulation of leptin expression and function in A/J vs. C57BL/6J mice during diet-induced obesity, Am. J. Physiol. Endocrinol. Metab., 279, E356–E365, 2000. Bergen, H.T., Mizuno, T., Taylor, J. and Mobbs, C.V., Resistance to dietinduced obesity is associated with increased proopiomelanocortin mRNA and decreased neuropeptide Y mRNA in the hypothalamus, Brain Res. 851, 198–203, 1999. Surwit, R.S., Edwards, C.L., Murthy, S. and Petro, A.E., Transient effects of long-term leptin supplementation in the prevention of diet-induced obesity in mice, Diabetes 49, 1203–1208, 2000. Zhang, J., Zhao, B., Gesongluobu, Sun, Y., Wu, Y., Pei, W., Ye, J., Hui, R. and Lui, L., Angiotensin-converting enzyme gene insertion/deletion (I/D) polymorphism in hypertensive patients with different degrees of obstructive sleep apnea, Hypertens. Res. 23, 407–411, 2000. Xiao, Y., Huang, X., Qiu, C., Zhu, X. and Liu, Y., Angiotensin I-converting enzyme gene polymorphism in Chinese patients with obstructive sleep apnea syndrome, Chin. Med. J. (Engl.) 112, 701–704, 1999. Barcelo, A., Elorza, M.A., Barbe, F., Santos, C., Mayoralas, L.R. and Agusti, A.G., Angiotensin converting enzyme in patients with sleep apnoea syndrome: plasma activity and gene polymorphisms, Eur. Respir. J. 17, 728–732, 2001. Agusti, A.G., Barbe, F. and Togores, B., Exhaled nitric oxide in patients with sleep apnea, Sleep 22, 231–235, 1999. Ip, M.S., Lam, B., Chan, L.Y., Zheng, L., Tsang, K.W., Fung, P.C. and Lam, W.K., Circulating nitric oxide is suppressed in obstructive sleep apnea and is reversed by nasal continuous positive airway pressure, Am. J. Respir. Crit. Care Med. 162, 2166–2171, 2000.
286 45.
46.
47.
48.
49.
50. 51.
52. 53. 54.
55. 56. 57.
58. 59. 60.
61.
Strohl Schulz, R., Schmidt, D., Blum, A., Lopes-Ribeiro, X., Lucke, C., Mayer, K., Olschewski, H., Seeger, W. and Grimminger, F., Decreased plasma levels of nitric oxide derivatives in obstructive sleep apnoea: response to CPAP therapy, Thorax 55, 1046–1051, 2000. Olopade, C.O., Christon, J.A., Zakkar, M., Hua, C., Swedler, W.I., Scheff, P.A. and Rubinstein, I., Exhaled pentane and nitric oxide levels in patients with obstructive sleep apnea, Chest 111, 1500–1504, 1997. Beall, C.M., Laskowski, D., Strohl, K.P., Soria, R., Villena, M., Vargas, E., Alarcon, A.M., Gonzales C. and Erzurum, E.C., Pulmonary nitric oxide in mountain dwellers, Nature 414, 411–412, 2001. Kato, M., Roberts-Thomson, P., Phillips, B.G., Haynes, W.G., Winnicki, M., Accurso, V. and Somers, V.K., Impairment of endothelium-dependent vasodilation of resistance vessels in patients with obstructive sleep apnea, Circulation 102, 2607–2610, 2000. Tahawi, Z., Orolinova, N., Joshua, I.G., Bader, M. and Fletcher, E.C., Altered vascular reactivity in arterioles of chronic intermittent hypoxic rats, J. Appl. Physiol. 90. 2007–2013, 2001 (discussion 2000). Lander, E.S. and Botstein, D., Mapping Mendelian factors underlying quantitative traits using RFLP linkage maps, Genetics 121, 185–199, 1989. Nadeau, J.H., Singer, J.B., Matin, A. and Lander, E.S., Analysing complex genetic traits with chromosome substitution strains, Nat. Genet. 24, 221–225, 2000. Shea, S.A., Life without ventilatory chemosensitivity, Respir. Physiol. 110, 199– 210, 1997. Richter, D.W. and Spyer, K.M., Studying rhythmogenesis of breathing: comparison of in vivo and in vitro models, Trends Neurosci. 24, 464–472, 2001. DeLorme, M.P. and Moss, O.R., Pulmonary function assessment by wholebody plethysmography in restrained versus unrestrained mice, J. Pharmacol. Toxicol. Methods 47, 1–10, 2002. Bartlett, D., Jr. and Tenney, S.M., Control of breathing in experimental anemia, Respir. Physiol. 10, 384–395, 1970. Epstein, M.A. and Epstein, R.A., A theoretical analysis of the barometric method for measurement of tidal volume, Respir. Physiol. 32, 105–120, 1978. Chaui-Berlinck, J.G. and Bicudo, J.E., The signal in total-body plethysmography: errors due to adiabatic–isothermic difference, Respir. Physiol. 113, 259– 270, 1998. Jacky, J.P., A plethysmograph for long-term measurements of ventilation in unrestrained animals, J. Appl. Physiol. 45, 644–647, 1978. Jacky, J.P., Barometric measurement of tidal volume: effects of pattern and nasal temperature, J. Appl. Physiol. 49, 319–325, 1980. Enhorning, G., van Schaik, S., Lundgren, C. and Vargas, I., Whole-body plethysmography, does it measure tidal volume of small animals? Can. J. Physiol. Pharmacol. 76, 945–951, 1998. Hamelmann, E., Schwarze, J., Takeda, K., Oshiba, A., Larsen, G.L., Irvin, C.G. and Gelfand, E.W., Noninvasive measurement of airway responsiveness in allergic mice using barometric plethysmography, Am. J. Respir. Crit. Care Med. 156, 766–775, 1997 [see comments].
Inheritance and Ventilatory Behavior in Animal Models 62.
63. 64.
65.
66. 67. 68.
69.
70.
71.
72.
73.
74. 75.
76. 77.
78. 79.
287
Mortola, J.P. and Frappell, P.B., On the barometric method for measurements of ventilation, and its use in small animals, Can. J. Physiol. Pharmacol. 76, 937– 944, 1998. Fenelon, K., Seifert, E.L. and Mortola, J.P., Hypoxic depression of circadian oscillations in sino-aortic denervated rats, Respir. Physiol. 122, 61–69, 2000. Tankersley, C.G., Irizarry, R., Flanders, S. and Rabold, R., Circadian rhythm variation in activity, body temperature, and heart rate between C3H/HeJ and C57BL/6J inbred strains, J. Appl. Physiol. 92, 870–877, 2002. Petak, F., Habre, W., Donati, Y.R., Hantos, Z. and Barazzone-Argiroffo, C., Hyperoxia-induced changes in mouse lung mechanics: forced oscillations vs. barometric plethysmography, J. Appl. Physiol. 90, 2221–2230, 2001. Kline, D.D. and Prabhakar, N.R., Peripheral chemosensitivity in mutant mice deficient in nitric oxide synthase, Adv. Exp. Med. Biol. 475, 571–579, 2000. Crabbe, J.C., Wahlsten, D. and Dudek, B.C., Genetics of mouse behavior: interactions with laboratory environment, Science 284, 1670–1672, 1999. Flandre, T.D., Leroy, P.L. and Desmecht, D.J., Effect of somatic growth, strain, and sex on double-chamber plethysmographic respiratory function values in healthy mice, J. Appl. Physiol. 94, 1129–1136, 2003. Han, F., Subramanian, S., Dick, T.E., Dreshaj, I.A. and Strohl, K.P., Ventilatory behavior after hypoxia in C57BL/6J and A/J mice, J. Appl. Physiol. 91, 1962–1970, 2001. Tankersley, C.G., Fitzgerald, R.S. and Kleeberger, S.R., Differential control of ventilation among inbred strains of mice, Am. J. Physiol. 267, R1371–R1377, 1994. Tankersley, C.G., Fitzgerald, R.S., Levitt, R.C., Mitzner, W.A., Ewart, S.L. and Kleeberger, S.R., Genetic control of differential baseline breathing pattern, J. Appl. Physiol. 82, 874–881, 1997. Tagaito, Y., Polotsky, V.Y., Campen, M.J., Wilson, J.A., Balbir, A., Smith, P.L., Schwartz, A.R. and O’Donnell, C.P., A model of sleep-disordered breathing in the C57BL/6J mouse, J. Appl. Physiol. 91, 2758–2766, 2001. Tankersley, C.G., Fitzgerald, R.S., Mitzner, W.A. and Kleeberger, S.R., Hypercapnic ventilatory responses in mice differentially susceptible to acute ozone exposure, J. Appl. Physiol. 75, 2613–2619, 1993. Tankersley, C.G., A genomic model for differential hypoxic ventilatory responses, Adv. Exp. Med. Biol. 475, 75–85, 2000. Tononi, G. and Cirelli, C., Modulation of brain gene expression during sleep and wakefulness: a review of recent findings, Neuropsychopharmacology 25, S28–S35, 2001. Han, F. and Strohl, K.P., Inheritance of ventilatory behavior in rodent models, Respir. Physiol. 121, 247–256, 2000. Chua, T.P., Clark, A.L., Amadi, A.A. and Coats, A.J., Relation between chemosensitivity and the ventilatory response to exercise in chronic heart failure, J. Am. Coll. Cardiol. 27, 650–657, 1996. Hutchison, A.A. and Olinsky, A., Hypoxic and hypercapnic response in asthmatic subjects with previous respiratory failure, Thorax 36, 759–763, 1981. Strohl, K. and Redline, S., State-of-the-art: recognition of obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 154, 279–289, 1996.
288 80.
81. 82.
83.
84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94. 95.
Strohl Kronenberg, R.S., Drage, C.W. and Stevenson, J.E., Acute respiratory failure and obesity with normal ventilatory response to carbon dioxide and absent hypoxic ventilatory drive, Am. J. Med. 62, 772–776, 1977. Hida, W., Role of ventilatory drive in asthma and chronic obstructive pulmonary disease, Curr. Opin. Pulm. Med. 5, 339–343, 1999. Weese-Mayer, D., Silvestri, J., Marazita, M. and Hoo, J., Congenital central hypoventilation syndrome: inheritance and relation to sudden infant death syndrome, Am. J. Med. Genet. 47, 360–367, 1993. Ou, L.C., Hill, N.S. and Tenney, S.M., Ventilatory responses and blood gases in susceptible and resistant rats to high altitude, Resp. Physiol. 58, 161–170, 1984. Ou, L.C. and Smith, R.P., Probable strain differences of rats in susceptibilities and cardio-pulmonary responses to chronic hypoxia, Respir. Physiol. 53, 367– 377, 1983. Connelly, C.A., Otto-Smith, M.R. and Feldman, J.L., Blockade of NMDA receptor-channels by MK-801 alters breathing in adult rats, Brain Res. 596, 99– 110, 1992. Tankersley, C.G., Elston, R.C. and Schnell, A.H., Genetic determinants of acute hypoxic ventilation: patterns of inheritance in mice, J. Appl. Physiol. 88, 2310–2318, 2000. Tankersley, C.G., Selected contribution: variation in acute hypoxic ventilatory response is linked to mouse chromosome 9, J. Appl. Physiol. 90, 1615–1622, 2001 (discussion 1606). Tankersley, C.G., DiSilvestre, D.A., Jedlicka, A.E., Wilkins, H.M. and Zhang, L., Differential inspiratory timing is genetically linked to mouse chromosome 3. J. Appl. Physiol. 85, 360–365, 1998. Tankersley, C.G., Rabold, R. and Mitzner, W., Differential lung mechanics are genetically determined in inbred murine strains, J. Appl. Physiol. 86, 1764–1769, 1999. Strohl, K.P., Thomas, A.J., St. Jean, P., Schlenker, E.H., Koletsky, R.J. and Schork, N.J., Ventilation and metabolism among rat strains, J. Appl. Physiol. 82, 317-323, 1997. Hodges, M.R., Forster, H.V., Papanek, P.E., Dwinell, M.R. and Hogan, G.E., Ventilatory phenotypes among four strains of adult rats, J. Appl. Physiol. 93, 974–983, 2002. Subramanian, S., Han, F., Erokwu, B.O., Dick, T.E. and Strohl, K.P., Do genetic factors influence the Dejours phenomenon? Adv. Exp. Med. Biol. 499, 209–214, 2001. Ahmed, M., Serrette, C., Kryger, M.H. and Anthonisen, N.R., Ventilatory instability in patients with congestive heart failure and nocturnal Cheyne– Stokes breathing, Sleep 17, 527–534, 1994. Lander, E. and Schork, N., The genetic dissection of complex traits, Science 265, 2037–2048, 1994. Strohl, K.P., Subramanian, S., Han, F., Principe, K. and Dick, T.E., Incorporating inheritance into models for understanding ventilatory behavior, Sleep Breath. 5, 47–51, 2001.
Inheritance and Ventilatory Behavior in Animal Models 96.
97.
98.
99.
100.
101.
102.
103.
104.
105.
106.
107.
108.
289
Kline, D.D., Yang, T., Premkumar, D.R., Thomas, A.J. and Prabhakar, N.R., Blunted respiratory responses to hypoxia in mutant mice deficient in nitric oxide synthase-3. J. Appl. Physiol. 88, 1496–1508, 2000. Kline, D.D., Yang, T., Huang, P.L. and Prabhakar, N.R., Altered respiratory responses to hypoxia in mutant mice deficient in neuronal nitric oxide synthase, J. Physiol. (Lond.) 511, 273–287, 1998. Kline, D.D., Overholt, J.L. and Prabhakar, N.R., Mutant mice deficient in NOS-1 exhibit attenuated long-term facilitation and short-term potentiation in breathing, J. Physiol. 539, 309–315, 2002. Renolleau, S., Dauger, S., Vardon, G., Levacher, B., Simonneau, M., Yanagisawa, M., Gaultier, C. and Gallego, J., Impaired ventilatory responses to hypoxia in mice deficient in endothelin-converting-enzyme-1, Pediatr. Res. 49, 705–712, 2001. Kuwaki, T., Cao, W.H., Kurihara, Y., Kurihara, H., Ling, G.Y., Onodera, M., Ju, K.H., Yazaki, Y. and Kumada, M., Impaired ventilatory responses to hypoxia and hypercapnia in mutant mice deficient in endothelin-1, Am. J. Physiol. 270, R1279–R1286, 1996. Huey, K.A., Low, M.J., Kelly, M.A., Juarez, R., Szewczak, J.M. and Powell, F.L., Ventilatory responses to acute and chronic hypoxia in mice: effects of dopamine D(2) receptors, J. Appl. Physiol. 89, 1142–1150, 2000. Grasemann, H., Lu, B., Jiao, A., Boudreau, J., Gerard, N.P. and De Sanctis, G.T., Targeted deletion of the neutral endopeptidase gene alters ventilatory responses to acute hypoxia in mice, J. Appl. Physiol. 87, 1266– 1271, 1999. Kline, D.D., Peng, Y.J., Manalo, D.J., Semenza, G.L. and Prabhakar, N.R., Defective carotid body function and impaired ventilatory responses to chronic hypoxia in mice partially deficient for hypoxia-inducible factor 1alpha, Proc. Natl. Acad. Sci. USA 99, 821–826, 2002. Nakamura, A., Kuwaki, T., Kuriyama, T., Yanagisawa, M. and Fukuda, Y., Normal ventilation and ventilatory responses to chemical stimuli in juvenile mutant mice deficient in endothelin-3, Respir. Physiol. 124, 1–9, 2001. Ptak, K., Burnet, H., Blanchi, B., Sieweke, M., De Felipe, C., Hunt, S.P., Monteau, R. and Hilaire, G., The murine neurokinin NK1 receptor gene contributes to the adult hypoxic facilitation of ventilation, Eur. J. Neurosci. 16, 2245–2252, 2002. Burnet, H., Bevengut, M., Chakri, F., Bou-Flores, C., Coulon, P., Gaytan, S., Pasaro, R. and Hilaire, G., Altered respiratory activity and respiratory regulations in adult monoamine oxidase A-deficient mice, J. Neurosci. 21, 5212–5221, 2001. Fu, X.W., Wang, D., Nurse, C.A., Dinauer, M.C. and Cutz, E., NADPH oxidase is an O2 sensor in airway chemoreceptors: evidence from Kþ current modulation in wild-type and oxidase-deficient mice. Proc. Natl. Acad. Sci. USA 97, 4374–4379, 2000. Thompson, R.J., Farragher, S.M., Cutz, E. and Nurse, C.A., Developmental regulation of O(2) sensing in neonatal adrenal chromaffin cells from wild-type and NADPH-oxidase-deficient mice, Pflu¨gers Arch. 444, 539–548, 2002.
290
Strohl
109. Archer, S.L., Reeve, H.L., Michelakis, E., Puttagunta, L., Waite, R., Nelson, D.P., Dinauer, M.C. and Weir, E.K., O2 sensing is preserved in mice lacking the gp91 phox subunit of NADPH oxidase, Proc. Natl. Acad. Sci. USA 96, 7944–7949, 1999. 110. He, L., Chen, J., Dinger, B., Sanders, K., Sundar, K., Hoidal, J. and Findone, S., Characteristics of carotid body chemosensitivity in NADPH oxidasedeficient mice, Am. J. Physiol. Cell Physiol. 282, C27–C33, 2002. 111. Roy, A., Rozanov, C., Mokashi, A., Daudu, P., Al-mehdi, A.B., Shams, H. and Lahiri, S., Mice lacking in gp91 phox subunit of NAD(P)H oxidase showed glomus cell [Ca(2þ)](i) and respiratory responses to hypoxia, Brain Res. 872, 188–193, 2000. 112. Dauger, S., Renolleau, S., Vardon, G., Nepote, V., Mas, C., Simonneau, M., Gaultier, C. and Gallego, J., Ventilatory responses to hypercapnia and hypoxia in Mash-1 heterozygous newborn and adult mice. Pediatr. Res. 46, 535–542, 1999. 113. Tankersley, C.G., Haxhiu, M.A. and Gauda, E.B., Differential CO(2)-induced c-fos gene expression in the nucleus tractus solitarii of inbred mouse strains, J. Appl. Physiol. 92, 1277–1284, 2002. 114. Nattie, E.E., Central chemosensitivity, sleep, and wakefulness. Respir. Physiol. 129, 257–268, 2001. 115. Tankersley, C.G., O’Donnell, C., Daood, M.J., Watchko, J.F., Mitznerm, W., Schwartz, A. and Smith, P., Leptin attenuates respiratory complications associated with the obese phenotype, J. Appl. Physiol. 85, 2261–2269, 1998. 116. O’Donnell, C.P., Tankersley, C.G., Polotsky, V.P., Schwartz, A.R. and Smith, P.L., Leptin, obesity, and respiratory function, Respir. Physiol. 119, 163–170, 2000. 117. Kline, D.D. and Prabhakar, N.R., Role of nitric oxide in short-term potentiation and long-term facilitation: involvement of NO in breathing stability, Adv. Exp. Med. Biol. 499, 215–219, 2001. 118. Subramanian, S., Erokwu, B., Han, F., Dick, T.E. and Strohl, K.P., L-NAME differentially alters ventilatory behavior in Sprague-Dawley and Brown Norway rats, J. Appl. Physiol. 93, 984–989, 2002. 119. Tankersley, C., Kleeberger, S., Russ, B., Schwartz, A. and Smith, P., Modified control of breathing in genetically obese (ob/ob) mice, J. Appl. Physiol. 81, 716–723, 1996. 120. Tankersley, C.G., Genetic control of ventilation: what are we learning from murine models? Curr. Opin. Pulm. Med. 5, 344–348, 1999. 121. Tankersley, C.G., Kulaga, H. and Wang, M.M., Inspiratory timing differences and regulation of Gria2 gene variation: a candidate gene hypothesis, Adv. Exp. Med. Biol. 499, 477–482, 2001. 122. Stoll, M., Cowley, A.W., Jr., Tonellato, P.J., Greene, A.S., Kaldunski, M.L., Roman, R.J., Dumas, P., Schork, N.J., Wang, Z. and Jacob, H.J., A genomicsystems biology map for cardiovascular function, Science 294, 1723–1726, 2001. 123. Beall, C.M., Strohl, K.P., Blangero, J., Williams-Blangero, S., Almasy, L.A., Decker, M.J., Worthman, C.M., Goldstein, M.C., Vargas, E., Villena, M., Soria, R., Alarcon, A.M. and Gonzales, C., Ventilation and hypoxic
Inheritance and Ventilatory Behavior in Animal Models
124.
125.
126.
127.
128.
129.
130.
291
ventilatory response of Tibetan and Aymara high altitude natives, Am. J. Phys. Anthropol. 104, 427–447, 1997. Vela-Bueno, A., Kales, A., Soldatos, C.R., Dobladez-Blanco, B., CamposCastello, J., Espino-Hurtado, P. and Olivan-Palacios, J., Sleep in the PraderWilli syndrome, Arch. Neurol. 41, 294–296, 1984. Hertz, G., Cataletto, M., Feinsilver, S.H. and Angulo, M., Developmental trends of sleep-disordered breathing in Prader-Willi syndrome: the role of obesity, Am. J. Med. Genet. 56, 188–190, 1995. Manni, R., Politini, L., Nobili, L., Ferrillo, F., Livieri, C., Venslli, E., Biancheri, R., Martinetti, M. and Tartara, A., Hypersomnia in the PraderWilli syndrome: clinical-electrophysiological features and underlying factors, Clin. Neurophysiol. 112, 800–805, 2001. Schluter, B., Buschatz, D., Trowitzsch, E., Aksu, F. and Andler, W., Respiratory control in children with Prader-Willi syndrome, Eur. J. Pediatr. 156, 65–68, 1997. Davies, J.L., Kawaguchi, Y., Bennett, S.T., Copeman, J.B., Cordell, H.J., Pritchard, L.E., Reed, P.W., Gough, S.C., Jenkins, S.C., Palmer, S.M., Balfour, K.M., Rowe, B.R., Farrall, M., Barnett, A.H., Bain, S.C. and Todd, J.A., A genome-wide search for human type 1 diabetes susceptibility genes, Nature 371, 130–135, 1994. Iyengar, S.K., Stein, C.M., Russo, K., Erokwu, B.O. and Strohl, K.P., The fa leptin receptor mutation and the heritability of respiratory frequency in a Brown Norway and Zucker intercross, J. Appl. Physiol. 97, 811–820, 2004, e-pub March 19, 2004. Dauger, S., Guimiot, F., Renolleau, S., Levacher, B., Boda, B., Mas, C., Nepote, V., Simonneau, M., Gaultier, C. and Gallego, J., MASH-1/RET pathway involvement in development of brain stem control of respiratory frequency in newborn mice, Physiol. Genomics 7, 149–157, 2001.
Part II Pathophysiology
9 Congenital Central Hypoventilation Syndrome: Should We Rename It Congenital Autonomopathy?
DAVID GOZAL University of Louisville Louisville, Kentucky
I.
Introduction
There is no doubt that in the context of disorders of respiratory control, congenital central hypoventilation syndrome (CCHS) occupies a unique place in human physiology. This entity, which can be viewed as a true experiment of nature, has allowed for major discoveries about the roles played by chemosensitivity in the maintenance of gas homeostasis as a function of state. However, as always occurs in biology, things turn out to be more complicated than anticipated. As such, the conceptual frameworks that have been proposed to account for the manifestations of CCHS have evolved over recent years to incorporate novel findings derived from physiology, genetics, and medicine. This chapter will review such findings and delineate the evolution of the concept that CCHS, rather than represent a pure phenotype of absent central chemosensitivity, is rather an intrinsic disorder of the autonomic nervous system.
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Definition and diagnosis
CCHS (Ondine’s curse) is traditionally defined as the failure of automatic control of breathing [1–10]. The term ‘‘Ondine’s curse’’ was initially employed by Severinghaus and Mitchell to describe three adult patients who developed central hypoventilation following high cervical/brainstem surgery [11]. When awake and requested to breathe voluntarily, these patients would have no difficulty in performing such task, yet they required mechanical ventilatory support for severe central apnea while asleep [11]. In 1970, an infant with the typical clinical features corresponding to CCHS was reported [1]. In CCHS, ventilation is most profoundly affected during quiet/ non-rapid-eye-movement sleep (NREMS), a state during which automatic neural control is predominant [5]. Ventilatory patterns are also abnormal during active/rapid eye movement sleep (REMS) and during wakefulness, but to a lesser degree [1,3,5,6,12–14]. The severity of ventilatory dysfunction can range from relatively mild alveolar hypoventilation during sleep and almost adequate alveolar ventilation during wakefulness, to complete apnea during sleep with severe alveolar hypoventilation during waking. Other symptoms indicative of brainstem dysfunction may be present, but are not essential to make the diagnosis of CCHS. The clinical presentation of CCHS varies depending on the severity of the phenotype [8,9,15–17]. Most CCHS patients will manifest their disorder in the newborn period; they may be apneic at birth or develop apnea and cyanosis requiring resuscitation in the delivery room or nursery, such that often the concern for preceding perinatal asphyxia usually overrides the suspicion of CCHS. However, while perinatal asphyxia of a magnitude leading to severe respiratory depression is usually associated with significant additional neurological manifestations, CCHS infants will essentially display intact neurological function provided that they receive adequate ventilatory support. The inability to wean these babies from mechanical ventilatory support along with the observation that their respiratory disturbance appears worse while the baby is asleep usually prompts the diagnosis. It should be pointed out that over the first several months or years of life, there is an apparent improvement in the magnitude of the alveolar hypoventilation in CCHS infants. However, rather than reflect a change in the underlying disorder, these dynamic temporal changes most likely represent the normal maturation of the respiratory system. The vast proportion of CCHS infants will not be apneic at birth and will develop cyanosis, tachycardia, and diaphoresis when asleep, and can be mistaken for having a congenital cardiac defect or be confused with apparent lifethreatening events [8]. It is assumed that some SIDS cases may in fact represent unrecognized CCHS. Others have such mild hypoventilation that they can go unrecognized for several years and present with pulmonary hypertension and cor pulmonale. It is estimated that about 30% of CCHS
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patients will require mechanical ventilatory support 24 h/day with the remainder displaying near adequate ventilation and gas exchange during wakefulness. It should be stressed that the diagnosis of CCHS is one of exclusion, i.e., the etiology of hypoventilation is unknown. Therefore, all other known causes of hypoventilation need to be ruled out, and this may be particularly difficult during the newborn period. Obviously, ventilatory muscle weakness, cardiac disease, or any type of respiratory disease all need to be excluded. Magnetic Resonance Imaging (MRI) scans of the brain including the brainstem will usually rule out the presence of anatomic lesions, which are absent in CCHS. Multiple metabolic disorders can present as apnea and cyanosis in the infant, and therefore a metabolic screen should be conducted. Specific disorders such as Leigh’s disease, pyruvate dehydrogenase deficiency [18], and carnitine deficiency should be considered as well as congenital myopathies, diaphragmatic dysfunction, congenital myasthenia gravis, or Mobius syndrome [19]. Because asphyxia, infection, trauma, or tumors may coexist, the decisions about the contribution of these confounding factors to the clinical presentation may be arduous [15]. In general, the initial evaluation should include chest X-rays, fluoroscopic or ultrasound evaluation of the diaphragm, electrocardiogram, echocardiogram, 24 h Holter recording, meticulous neurologic examination in addition to the Central Nervous System (CNS) imaging studies, and urinary and plasma screen for metabolic disturbances. A detailed ophthalmological examination may reveal abnormal pupillary and optical disk features [20–22]. In those cases where mild abdominal distension or delayed defecation is present, a rectal biopsy should be considered to determine whether Hirschprung’s disease (HD) may be present, since it will occur in 15–20% of all patients with CCHS [14,22–30]. A detailed evaluation in a respiratory physiology laboratory is critically important to the diagnosis and management of these patients. Polygraphic recordings of respiratory and cardiac signals during all sleep and wake stages, including chest and abdominal wall motion (respiratory inductance plethysmography), end-tidal CO2, pulse oximetry and waveform, and ECG can be combined to EEG and EMG channels that will define state. Careful observation of the infant’s respiratory (tidal volume and frequency) and cardiac frequency responses to ongoing spontaneous changes in blood gas exchange can provide important information and obviate the need for immediate application of exogenous hypercapnic and hypoxic challenges. Such recordings will also permit determination of the appropriate levels of mechanical respiratory support that the patient needs during the various states. It should be emphasized that these patients can be very unstable, particularly during their early years of life. Indeed, minor respiratory infections can trigger apnea both during wakefulness and during sleep, and the absence of subjective or objective response to hypoxia or
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hypercapnia in these patients, which in normal children would manifest as increased respiratory efforts, retractions, or nasal flaring, will further mask the progression of potentially serious deterioration in clinical status. Thus, unless very subtle symptoms are accounted for, hypoxia for example will be detected only when lethargy, cyanosis, and/or loss of consciousness survene. Thus, continuous monitoring by trained and skilled observers is necessary to prevent significant hypoxemia and its sequelae. Similarly, bradycardia is not uncommon, even if it does not usually require implantation of cardiac pacemakers in infancy [9,31–35]. Syncope, particularly during Valsalva-like maneuvers, is not uncommon as well [36]. Feeding difficulties during infancy and abnormal gastroesophageal motility mostly presenting as gastroesophageal reflux are frequent and can lead to aspiration of intestinal contents, resulting in frequent implementation of gastrostomy and anti-reflux procedures [37–39]. In practical terms, the proposed diagnostic criteria for CCHS have been recently reviewed [15] and essentially consist of the following: 1. persistent evidence of sleep hypoventilation (PaCO2 4 60 mmHg); 2. the onset of symptoms usually occurred during the first year of life, and in most during the first few days of life; 3. absence of primary pulmonary disease or neuromuscular dysfunction, which could explain the hypoventilation; 4. no evidence of cardiac disease; 5. no evidence of metabolic disorders.
III.
Pathophysiology
The exact pathophysiology of CCHS remains unknown. However, most of the evidence now suggests that this is a generalized disorder of the autonomic nervous system which affects many more systems than just the control of respiration. Indeed, in addition to HD, multiple mediastinal and adrenal ganglioneuromas and other tumors of the neural crest have been described in CCHS patients [23,25,40–44]. Decreased heart rate variability is almost universal in CCHS [32], and with increasing age, arrhythmias may occur and may necessitate implantation of cardiac pacemakers [34]. Furthermore, specific testing of cardiac responses to baroreceptor stimulation in children with CCHS, revealed marked alteration in cardiovascular fast-component characteristics of the reflex [45]. The gastrointestinal dismotility that is so frequent in early life further supports the notion of a diffuse autonomic disorder. Ophthalmological abnormalities, especially those mediating the neural control of eye movement and pupillary responses, are frequently seen in CCHS patients [20–22]. In addition, brainstem auditory evoked potentials can be abnormal [46–48].
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Recent and exciting developments have occurred lending support to the genetic hypothesis of CCHS. Indeed, a genetic defect is the probable underlying etiology for many, if not all, CCHS patients, and this assumption is particularly enhanced in the context of CCHS coinciding with the presence of HD. While a single gene hypothesis has been proposed, it is more likely that CCHS is the phenotypic manifestation resulting from the interaction between the environment and a polygenic disease. The rationale for CCHS containing a genetic component is its early manifestation in the newborn period, and its occurrence in families including siblings, female twins, mother with neuroblastoma and child with CCHS, and more recently mothers with CCHS giving birth to infants with CCHS [31,49–53], and of course its association with HD [14,22–30]. In fact, up to 20% of reported cases of CCHS are accompanied by HD, such that the association of these two relatively rare clinical entities suggests a possible common pathogenetic basis. Screening for mutations in CCHS, however, has yielded informative yet somewhat disappointing results. Indeed, since a mutation in the RETprotooncogene is associated with HD [54–56], and since RET may play a critical role in the development of the neural crest and parasympathetic system in both HD [54] and CCHS [40–44], a genetic screening of RET mutations was performed by several research groups and revealed that occasional patients may exhibit RET mutations, even when HD is not present [54–61]. Similarly, mutations in the endothelin 3 gene, brain-derived neurotrophic factor [62–64], and in glial-derived neurotrophic factor [65] have been described in patients with CCHS. However, the significance of such sporadic findings remains to be established. In a major, recent breakthrough discovery in this field, mutations in the Phox2b gene, a gene critically involved in the development of the neural crest during embryogenesis, have been identified in a large proportion of French children with CCHS and their families [66]. These findings open new and exciting venues for the understanding of the pathophysiological mechanisms underlying the phenotypic manifestations of CCHS, and further provide conceptual support for the title of this chapter, especially considering the important role of neural crest-derived neurons in autonomic function. It should be emphasized that the putative genetic etiology of CCHS is at least partially undermined by the fact that family members of CCHS patients do not display any evidence of respiratory control dysfunction [67]. In contrast, a recent case/control study revealed that families and siblings of CCHS patients exhibit higher frequency of symptoms compatible with autonomic alterations, thereby suggesting that subtle phenotypic manifestations may be present and supporting some form of Mendelian inheritance [68]. Creation and implementation of a genomic/phenotype database of all diagnosed patients with CCHS will undoubtedly facilitate future genetic studies in this disorder.
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Animal Models
A true animal model displaying all or most of the phenotypic expression patterns of CCHS has not yet become available. Schla¨fke and colleagues have demonstrated that after either electrocoagulation- or ibotenic acidinduced lesions of the intermediate area within the ventral medullary surface, significant compromise of both hypoxic and hypercapnic ventilatory responses will occur in anesthetized and awake cats [69–71]. This experimental model shares some of the typical respiratory control alterations found in CCHS patients. In recent years, and despite the aforementioned negative genetic findings, several investigators have explored the possibility that genetic manipulation of the RET protooncogene in the mouse may lead to respiratory manifestations reminiscent of CCHS [72]. Indeed, markedly reduced ventilatory responses to hypercapnia were present in RET knockout mice [72]. Using whole-body flow plethysmography, baseline breathing and ventilatory and arousal responses to chemical stimuli were examined in unrestrained heterozygous c-ret þ/ newborn mice and their wild-type c-ret þ/þ litter mates at 10–12 h of postnatal age [73]. The hyperpneic and arousal responses to hypoxia and hypercapnia were not significantly different in these two groups. However, the number and total duration of apnea and periodic breathing episodes were significantly higher in c-ret þ/ than in c-ret þ/þ pups during hypoxia and post-hypoxic normoxia. Thus, different aspects of respiratory control are determined by the activity of neural crest controlling genes. Erickson et al. reported similar results in brain-derived growth factor knock-out mice [74], thereby suggesting that multiple genes may be involved in the regulation and development of respiratory and autonomic control sites during embryogenesis. In this regard, recent work with homeobox genes led to the generation of a knockout mouse for RNX. The genes Tlx1 (Hox11), Enx (Hox11L2, Tlx-2) and Rnx (Hox11L2, Tlx-3) constitute a family of orphan homeobox genes. In situ hybridization has revealed considerable overlap in their expression within the nervous system, but Rnx is singularly expressed in the developing dorsal and ventral region of the medulla oblongata. Transgenic mice in which the RNX gene was knocked-out revealed severe hypoventilation, suggesting that this gene may be implicated in CCHS [75]. However, three recent screens for mutations in the RNX gene failed to reveal any abnormality in CCHS patients [60,76,77]. V.
Structural Central Nervous System Abnormalities
Based on the initial assumption that a centrally located defect would be present in CCHS, many attempts have been made over the years to identify
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structural CNS abnormalities. Earlier reports of hypoplasia of the arcuate nucleus in one patient with CCHS [78], and the presence of abnormal evoked potential responses to auditory stimuli [46–48] further suggested that a brainstem lesion might be present. In addition, central hypoventilation syndrome has been described in occasional patients with cerebrovascular malformations and in patients with CNS infections [24,79–83]. Careful radiological surveys of the brain in several CCHS patients have thus far failed to identify any recognizable lesion accountable for the unique manifestations of this syndrome [84] (Harper, personal communication). Neuronal loss in reticular nuclei and cranial nerve nuclei as well as in the nucleus ambiguus and the hypoglossal and dorsal motor nucleus of the vagus have been reported in an infant with CCHS [81]. Furthermore, Cutz et al. reported that in two CCHS patients, their carotid bodies had volumes less than 50% of those seen in normal individuals with reduced numbers of glomus cells [85]. These investigators also reported substantial hypertrophy in the neuroepithelial bodies of these CCHS patients, possibly as a compensatory mechanism for the carotid body abnormality [85]. More recently, using non-invasive functional MRI approaches which provide functional topographic maps of the brain in response to the application of specific stimulation paradigms [86–90] significantly reduced responses in several brain regions in CCHS patients further suggesting that the ‘‘defect’’ is diffusely represented in regions that are immediately pertinent to the embryogenesis of the neural crest [91–96]. However, such a diffuse pattern of neural recruitment deficits also reinforces the concept that the global defect may involve integration of neural autonomic afferent inputs rather than represent a defect in intrinsic chemosensitivity. VI.
Physiologic Abnormalities of Ventilatory Control
Identification of the putative site(s) underlying the phenotypic manifestations of CCHS could not only lead to better therapeutic approaches in these patients, but also provide extremely important insights into modeling and localization of structures mediating respiratory control. Dissection of each of the components of the various functional elements involved in CCHS is obviously impossible in humans. Within these limitations, the following studies provide important information regarding the behavioral characteristics of respiratory control systems in CCHS. A schematic diagram of such approach is provided in Figure 9.1, and should assist in the rationale and interpretation of the studies conducted below. The initial corollary examined was whether voluntary breathing could be affected. Children with CCHS demonstrate no deficit in generating volitional breathing. The observation that the magnitude of alveolar hypoventilation was principally expressed during NREMS led to
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Arousal centers Cortex
CENTRAL CHEMORECEPTORS
(2) (1) ???
Hypothalamus locomotion center
Central respiratory drive (4)
Respiratory muscles (3)
Spinal relays (5) Mechanoreceptors muscle afferents
Pulmonary venous return
· VE Peripheral chemoreceptors PaO , PaCO , pH 2
2
Figure 9.1 Schematic diagram of a simplified model of the respiratory control system with potential chemoreceptive, mechanoreceptive and behavioral inputs to the medullary respiratory controller (pathways 1–5), and to the reticular activating system (pathway 6). In CCHS subjects, cortical pathways (1) are operational. The central chemoreceptive pathway and/or putative integrative chemoreceptive pathway (2) are defective. However, pathway 3 may be partially functional as abrupt peripheral chemoreceptor stimulation may induce some ventilatory changes in CCHS subjects. The hypothalamic/locomotor center (4) and mechanoreceptive pathway (5) have been demonstrated to effectively stimulate ventilation in CCHS. For discussion, see text.
the assumption that central chemoreceptor function was abnormally reduced or absent. Examination of ventilatory responses to endogenous challenges of isolated hypercapnia, hypoxemia, and to combined hypoxia and hypercapnia (asphyxia) revealed negligible responses during sleep [1–5,8,9], as well as during wakefulness [97]. Indeed, Paton and coworkers studied rebreathing hypoxic and hypercapnic ventilatory responses during wakefulness in five children with CCHS aged 6–11 years, and found that ventilatory responses to both hypoxia and hypercapnia were essentially random, with no evidence of progressive ventilatory increases despite increasing stimulus. Interestingly, despite absent rebreathing ventilatory responses to both hypercapnia and hypoxia [97], most CCHS patients are able to sustain adequate ventilation during wakefulness [98], such that redundant mechanisms may be operational. A recent study by Gaultier et al. demonstrated that while alveolar hypoventilation was particularly
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prominent during NREM sleep, hypercapnic chemosensitivity was unaffected by sleep states [99]. These data suggest that the intrinsic defect in CCHS is present at all times, but becomes more prominently expressed during conditions in which other redundant mechanisms are either less active or inoperative. One such mechanism could be related to either intact or residual peripheral chemoreceptor function. In five children with CCHS (ages 9–14 years) who demonstrated adequate ventilation during wakefulness, ventilatory challenges with 100% oxygen breathing, five tidal breaths of 100% N2, and vital capacity breaths of 5% and 15% CO2 in O2 and 5% CO2 in N2 induced overall similar increases in minute ventilation (VE) in CCHS and controls [100], suggesting that during abrupt transients in inspired gas concentrations, the peripheral chemoreceptors can be stimulated and induce intact responses [100]. The greater interindividual variability of responses in CCHS could reflect the lack of modulation of ventilatory control due to defective integration of afferent respiratory neural input [100,101]. In this context, if indeed the deficit in CCHS corresponds to defective integration of autonomic sensory stimuli, then arousal during hypercapnia should occur even if the ventilatory responses to hypercapnia are absent. Indeed, the frequency of arousal from a hypercapnic challenge during sleep was similar in eight CCHS and eight controls [102], suggesting the presence of intact central chemoreceptor sensitivity, and of defective integration of chemoreceptor inputs [102]. Based on the premise that chemoreceptors are important controllers of ventilation during exercise, one would also expect to observe substantial gas exchange alterations during physical activity in CCHS patients. In fact, Silvestri et al. showed severe gas exchange abnormalities in full-time ventilator-dependent CCHS patients during moderate exercise [103]. However, both Paton et al. [104] and Shea et al. [105] showed that exercise-induced hyperpnea can occur in CCHS patients who require ventilatory assistance during sleep, particularly at exercise intensities below the anaerobic threshold [103–105]. Such observations suggest that in the absence of ventilatory response to gradual chemoreceptor stimulation, movement exerts a dominant influence on respiratory rate, and consequently on the increase of VE during exercise [104,105], further strengthening the concept that central integration of chemosensory and metabolic inputs is faulty in CCHS. To further examine this concept, passive lower extremity motion approaches were used, both during wakefulness and sleep. Passive leg motion elicited relative hyperventilation in excess of metabolic requirements, resulting in normalization of end-tidal carbon dioxide tension (PETCO2 ), independent of state [106,107]. Thus, in a setting of deficient integration of respiratory control inputs, either mechanoreceptor afferent input from muscle and joints and/or rhythmic entrainment of respiration takes over, and plays a significant role in the modulation of breathing
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during exercise in children with CCHS [106–108]. Furthermore, normalization of PETCO2 with motion as measured in CCHS would lend support to a basic defect in integration of efferent and afferent neural inputs to respiratory controllers sites. VII.
Autonomic Nervous System Dysfunction
In addition to disturbed moment-to-moment heart rate variability [32–35], significant alterations in dopamine turnover have been found in patients with CCHS [109], and vagally mediated syncope may also occur [36], thereby lending further support to the assumption that significant dysregulation of central autonomic nervous system control is frequently present in CCHS. As mentioned, such alterations are also present in family members, and segregation analysis further indicates that CCHS is the most severe manifestation of a generalized dysfunction of the autonomic nervous system with a family pattern consistent with Mendelian transmission [110]. VIII.
Summary and Conclusions
The clinical and physiological spectrum of CCHS supports major physiological concepts, namely, (a) this disorder represents a congenital and genetically determined disease variant of the autonomic nervous system in general, and more specifically of the embryological development of the neural crest, and (b) the recruitment and activation of redundant mechanisms in these patients allows for the ability to sustain near adequate gas exchange during wakefulness and physical activity. Further efforts to expand on genotype–phenotype relationships and interactions should permit better insights into the normal development and function of the autonomic nervous system in man. Acknowledgment The author is supported by grants from the National Institutes of Health (HL69932, HL63912, and HL66358), and The Commonwealth of Kentucky Research Challenge Trust Fund. References 1.
Mellins, R.B., Balfour, H.H., Jr., Turino, G.M. and Winters, R.W., Failure of automatic control of ventilation (Ondine’s curse), Medicine 49, 487–504, 1970. 2. Fishman, L.S., Samson, J.H. and Sperling, D.R., Primary alveolar hypoventilation syndrome (Ondine’s curse), Am. J. Dis. Child 110, 155–161, 1965.
Congenital Central Hypoventilation Syndrome 3. 4.
5. 6.
7. 8.
9.
10.
11. 12.
13. 14.
15.
16.
17. 18.
305
Deonna, T., Arczynska, W. and Torrado, A., Congenital failure of automatic ventilation (Ondine’s curse), J. Pediatr. 84, 710–714, 1974. Shannon, D.C., Marsland, D.W., Gould, J.B., Callahan, B., Todres, I.D. and Dennis, J., Central hypoventilation during quiet sleep in two infants, Pediatrics 57, 342–346, 1976. Fleming, P.J., Cade, D., Bryan, M.H. and Bryan, A.C., Congenital central hypoventilation and sleep state, Pediatrics 66, 425–428, 1980. Guilleminault, C., McQuitty, J., Ariagno, R.L., Challamel, M.J., Korobkin, R. and McClead, R.E., Congenital central alveolar hypoventilation syndrome in six infants, Pediatrics 70, 684–694, 1982. Oren, J., Kelly, D.H. and Shannon, D.C., Long-term follow-up of children with congenital central hypoventilation syndrome, Pediatrics 80, 375–380, 1987. Marcus, C.L., Jansen, M.T., Poulsen, M.K., Keens, S.E., Nield, T.A., Lipsker, L.E. and Keens, T.G., Medical and psychosocial outcome of children with congenital central hypoventilation syndrome, J. Pediatr. 119, 888–895, 1991. Weese-Mayer, D.E., Silvestri, J.M., Menzies, L.J., Morrow-Kenny, A.S., Hunt, C.E. and Hauptman, S.A., Congenital central hypoventilation syndrome: diagnosis, management, and long-term outcome in thirty-two children, J. Pediatr. 120, 381–387, 1992. Keens, T.G. and Hoppenbrouwers, T., Congenital central hypoventilation syndrome (770.81), in Diagnostic Classification Steering Committee of the American Sleep Disorders Association, The International Classification of Sleep Disorders: Diagnostic and Coding Manual, Lawrence, KS, Allen Press, pp. 205–209, 1990. Severinghaus, J.W., and Mitchell, R.A., Ondine’s curse: failure of respiratory center automaticity while awake, Clin. Res. 10, 122, 1962. Hunt, C.E., Matalon, S.V. and Thompson, T.T., Central hypoventilation syndrome: experience with bilateral phrenic nerve pacing in 3 neonates, Am. Rev. Respir. Dis. 118, 23–28, 1978. Wells, H.H., Kattwinkel, J. and Morrow, J.D., Control of ventilation in Ondine’s curse. J. Pediatr. 96, 865–867, 1980. Haddad, G.G., Mazza, N.M., Defendini, R., Blanc, W.A., Driscoll, J.M., Epstein, A.F., Epstein, R.A. and Mellins, R.B., Congenital failure of automatic control of ventilation, gastrointestinal motility and heart rate, Medicine 57, 517–526, 1978. American Thoracic Society Consensus Statement, Idiopathic congenital central hypoventilation syndrome: diagnosis and management, Am. J. Respir. Crit. Care Med. 160, 368–373, 1999. Keens, T.G. and Davidson Ward, S.L. Syndromes affecting respiratory control during sleep, in Sleep and Breathing in Children – A Developmental Approach. Lung Biology in Health and Disease, Loughlin, G.M., Marcus, C.L. and Carrol, J.L., eds., New York, Marcel Dekker, Inc., pp. 525–553, 2000. Gozal, D., Congenital central hypoventilation syndrome: an update. Pediatr. Pulmonol. 26, 273–282, 1998. Johnston, K., Newth, C.J., Sheu, K.F., Patel, M.S., Heldt, G.P., Schmidt, K.A. and Packman, S., Central hypoventilation syndrome in pyruvate dehydrogenase complex deficiency, Pediatrics 74, 1034–1040, 1984.
306 19.
20. 21.
22.
23.
24.
25.
26.
27.
28.
29. 30.
31.
32.
33.
34.
Gozal Nunes, M.L., Friedrich, M.A. and Loch, L.F., Association of misoprostol, Moebius syndrome and congenital alveolar hypoventilation syndrome: a case report, Arq. Neuropsiquiatr. 57, 88–91, 1999. Dooling, E.C. and Richardson, E.P. Jr., Ophthalmoplegia and Ondine’s curse, Arch. Ophthalmol. 95, 1790–1793, 1977. Goldberg, D.S. and Ludwig, I.H., Congenital central hypoventilation syndrome: ocular findings in 37 children, J. Pediatr. Ophthalmol. Strabism. 33, 176–181, 1996. Lambert, S.R., Yang, L.L. and Stone, C., Tonic pupil associated with congenital neuroblastoma, Hirschsprung disease and central hypoventilation syndrome, Am. J. Opthalmol. 130, 238–240, 2000. Roshkow, J.E., Haller, J.A., Berdon, W.E. and Sane, S.M., Hirschsprung’s disease, Ondine’s curse, and neuroblastoma—manifestations of neurocristopathy, Pediatr. Radiol. 19, 45–49, 1988. Mukhopadhyay, S. and Wilkinson, P.W. Cerebral arteriovenous malformation, Ondine’s curse, and Hirschsprung disease, Dev. Med. Chil. Neurol. 32, 1087–1089, 1990. Stovroff, M., Dykes, F. and Teague, W.G., The complete spectrum of neurocristopathy in an infant with congenital hypoventilation, Hirschsprung’s disease, and neuroblastoma, J. Pediatr. Surg. 30, 1218–1221, 1995. Minutillo, C., Pemberton, P.J. and Goldblatt, J., Hirschsprung’s disease and Ondine’s curse: further evidence for a distinct syndrome, Clin. Genet. 36, 200–203, 1989. Nakahara, S., Yokomori, K., Tamura, K., Oku, K., Tsuchida, Y. and Hirschsprung’s disease associated with Ondine’s curse: a special subgroup? J. Pediatr. Surg. 10, 1481–1484, 1995. el-Halaby, E. and Coran, A.G., Hirschsprung’s disease associated with Ondine’s curse: report of three cases and review of the literature, J. Pediatr. Surg. 29, 530–535, 1994. Fodstad, H., Ljunggren, B. and Shawis, R., Ondine’s curse with Hirschsprung’s disease, Br. J. Neurosurg. 4, 87–93, 1990. Verloes, A., Elmer, C., Lacombe, D., Heinrichs, C., Rebuffat, E., Demarquez, J.L., Moncla, A. and Adam, E., Ondine-Hirschsprung syndrome (Haddad syndrome). Further delineation in two cases and review of the literature, Eur. J. Pediatr. 152, 75–77, 1993. Hamilton, J. and Bodurtha, J.N., Congenital central hypoventilation syndrome and Hirschsprung’s disease in half sibs, J. Med. Genet. 26, 272–274, 1989. Woo, M.S., Woo, M.A., Gozal, D., Jansen, M.T., Keens, T.G. and Harper, R.M., Heart rate variability in congenital central hypoventilation syndrome, Pediatr. Res. 31, 291–296, 1992. Ogawa, T., Kojo, M., Fukushima, N., Sonoda, H., Goto, K., Ishiwa, S., Ishiguro, M., Cardio-respiratory control in an infant with Ondine’s curse: a multivariate autoregressive modelling approach, J. Auton. Nerv. Syst., 42, 41–52, 1993. Silvestri, J.M., Hanna, B.D., Volgman, A.S., Jones, P.J., Barnes, S.D. and Weese-Mayer, D.E., Cardiac rhythm disturbances among children with
Congenital Central Hypoventilation Syndrome
35. 36.
37.
38.
39.
40.
41.
42. 43. 44.
45.
46. 47.
48. 49.
50.
307
idiopathic congenital central hypoventilation syndrome, Pediatr. Pulmonol. 29, 351–358. 2000. Lang, U., Braems, G. and Kunzel, W., Heart rate alterations in a fetus with Ondine’s curse, Gynecol. Obstet. Invest. 30, 124–126, 1990. O’Sullivan, J., Cottrell, A.J. and Wren, C., Ondine’s curse and neurally mediated syncope—a new and important association, Eur. Heart J. 14, 1289–1291, 1993. Faure, C., Viarme, F., Cargill, G., Navarro, J., Gaultier, C. and Trang, H., Abnormal esophageal motility in children with congenital central hypoventilation syndrome, Gastroenterology 122, 1258–1263, 2002. Alvord, E.C. Jr. and Shaw, C.M., Congenital difficulties with swallowing and breathing associated with maternal polyhydramnios: neurocristopathy or medullary infarction? J. Child. Neurol. 4, 299–306, 1989. Takeda, S., Fujii, Y., Kawahara, H., Nakahara, K. and Matsuda, H., Central alveolar hypoventilation (Ondine’s curse) with gastroesophageal reflux, Chest 110, 850–852, 1996. Rohrer, T., Trachsel, D., Engelcke, G. and Hammer, J., Congenital central hypoventilation syndrome associated with Hirschsprung’s disease and neuroblastoma: case of multiple neurocristopathies, Pediatr. Pulmonol. 33, 71–76, 2002. Swaminathan, S., Gilsanz, V., Atkinson, J. and Keens, T.G., Congenital central hypoventilation syndrome associated with multiple ganglioneuromas, Chest 96, 423–424, 1989. Poceta, J.S., Strandjord, T.P., Badura, R.J., Jr. and Milstein, J.M., Ondine curse and neurocristopathy, Pediatr. Neurol. 6, 370–372, 1987. Gaisie, G., Hirschsprung’s disease, Ondine’s curse, and neuroblastoma— manifestations of neurocristopathy, Pediatr. Neurol. 3, 370–372, 1987. Diez Garcia, R., Carrillo, A., Bartolome, M., Casanova, A. and Prieto, M., Central hypoventilation syndrome associated with ganglioneuroblastoma, Eur. J. Pediatr. Surg. 5, 292–294, 1995. Kim, A.H., Macey, P.M., Woo, M.A., Yu, P.L., Keens, T.G., Gozal, D. and Harper, R.M., Cardiac responses to pressor challenges in congenital central hypoventilation syndrome, Somnologie, 6, 109–115, 2002. Long, K.J. and Allen, N., Abnormal brain-stem auditory evoked potentials following Ondine’s curse, Arch. Neurol. 41, 1109–1110, 1984. Litscher, G., Schwarz, G. and Reimann, R., Abnormal brain stem auditory evoked potentials in a girl with the central alveolar hypoventilation syndrome, Int. J. Neurosci. 87, 113–117, 1996. Beckerman, R.C., Meltzer, J., Sola, A., Dunn, D. and Weggman, M., Brainstem auditory response in Ondine’s syndrome, Arch. Neurol. 43, 698–701, 1986. Kerbl, R., Litscher, H., Grubbauer, H.M., Reiterer, F., Zoble, G., Trop, M., Urlesberger, B., Eber, E. and Kurz, R., Congenital central hypoventilation syndrome (Ondine’s curse syndrome) in two siblings: delayed diagnosis and successful noninvasive treatment, Eur. J. Pediatr. 155, 977–980, 1996. Weese-Mayer, D.E., Silvestri, J.M., Marazita, M.L. and Hoo, J.J., Congenital central hypoventilation syndrome: inheritance and relation to sudden infant death syndrome, Am. J. Med. Genet. 47, 360–367, 1993.
308 51. 52.
53.
54.
55.
56.
57.
58.
59. 60.
61.
62.
63.
64.
Gozal Khalifa, M.M., Flavin, M.A. and Wherrett, B.A., Congenital central hypoventilation syndrome in monozygotic twins, J. Pediatr. 113, 853–855, 1988. Sritippayawan, S., Hamutcu, R., Kun, S.S., Ner, Z., Ponce, M. and Keens, T.G., Mother–daughter transmission of congenital central hypoventilation syndrome, Am. J. Respir. Crit. Care Med. 166, 367–369, 2002. Silvestri, J.M., Chen, M.L., Weese-Mayer, D.E., McQuitty, J.M., Carveth, H.J., Nielson, D.W., Borowitz, D. and Cerny, F., Idiopathic congenital central hypoventilation syndrome: the next generation, Am. J. Med. Genet. 112, 46–50, 2002. Edery, P., Lyonnet, S., Mulligan, L.M., Pelet, A., Dow, E., Abel, L., Holder, S., Nihoul-Fekete, C., Ponder, B.A. and Munnich, A., Mutations of the RET proto-oncogene in Hirschsprung’s disease, Nature 367, 378–380, 1994. Romeo, G., Ronchetto, P., Luo, Y., Barone, V., Seri, M., Ceccherini, I., Pasini, B., Bocciardi, R., Lerone, M. and Kaariainen, H., Point mutations affecting the tyrosine kinase domain of the RET proto-oncogene in Hirschsprung’s disease, Nature 367, 377–378, 1994. Schuchardt, A., D’Agati, V., Larsson-Blomberg, L., Costantini, F. and Pachnis, V., Defects in the kidney and enteric nervous system of mice lacking the tyrosine kinase receptor Ret, Nature 367, 380–383, 1994. Amiel, J., Attie, T., Simeoni, J., Edery, P., Gaultier, C., Munnich, A. and Lyonnet, S., Mutation of the ret proto-oncogene in a patient with congenital central hypoventilation syndrome (Ondine’s curse) and Hirschsprung disease, Am. J. Hum. Gen. 57, A205, 1995. Bolk, S., Angrist, M., Schwartz, S., Silvestri, J.M., Weese-Mayer, D.E. and Chakravarti, A., Congenital central hypoventilation syndrome: mutation analysis of the receptor tyrosine kinase RET, Am. J. Hum. Gen. 63, 603–609, 1996. Kinane, T.B. and Burton, M.D., A genetic approach to the congenital central hypoventilation syndrome, Pediatr. Pulmonol. 23, 133–135, 1997. Kanai, M., Numakura, C., Sasaki, A., Shirahata, E., Akaba, K., Hashimoto, M., Hasegawa, H., Shirasawa, S. and Hayasaka, K., Congenital central hypoventilation syndrome: a novel mutation of the RET gene in an isolated case. Tohoku J. Exp. Med. 196, 241–246, 2002. Sakai, T., Wakizaka, A., Matsuda, H., Nirasawa, Y. and Itoh, Y., Point mutation in exon 12 of the receptor tyrosine kinase proto-oncogene RET in Ondine-Hirschsprung syndrome, Pediatrics 101, 924–926, 1998. Bolk, S., Angrist, M., Xie, J., Yanagisawa, M., Silvestri, J.M., Weese-Mayer, D.E. and Chakravarti, A., Endothelin-3 frameshift mutation in congenital central hypoventilation syndrome (letter), Nat. Genet. 13, 395–396, 1996. Sakai, T., Wakizaka, A. and Nirasawa, Y., Congenital central hypoventilation syndrome associated with Hirschsprung’s disease: mutation analysis of the RET and endothelin-signaling pathways, Eur. J. Pediatr. Surg. 11, 335–337, 2001. Weese-Mayer, D.E., Bolk, S., Silvestri, J.M. and Chakravarti, A., Idiopathic congenital central hypoventilation syndrome: evaluation of brain-derived neurotrophic factor genomic DNA sequence variation, Am. J. Med. Genet. 107, 306–310, 2002.
Congenital Central Hypoventilation Syndrome 65.
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
309
Amiel, J., Salomon, R., Attie, T., Pelet, A., Trang, H., Mokhtari, M., Gaultier, C., Munnich, A. and Lyonnet, S., Mutations of the RET-GDNF signaling pathway in Ondine’s curse, Am. J. Hum. Genet. 62, 715–717, 1998. Amiel, J., Laudier, B., Attie-Bitach, T., Trang, H., de Pontual, L., Gener, B., Trochet, D., Etchevers, H., Ray, P., Simonneau, M., Vekemans, M., Munnich, A., Gaultier, C. and Lyonnet, S., Polyalanine expansion and frameshift mutations of the paired-like homeobox gene PHOX2B in congenital central hypoventilation syndrome, Nat. Genet. 33, 459–461, 2003. Marcus, C.L., Livingston, F.R., Wood, S.E. and Keens, T.G., Hypercapnic and hypoxic ventilatory responses in parents and siblings of children with congenital central hypoventilation syndrome, Am. Rev. Respir. Dis. 144, 136–140, 1991. Weese-Mayer, D.E., Silvestri, J.M., Huffman, A.D., Smok-Pearsall, S.M., Kowal, M.H., Maher, B.S., Cooper, M.E. and Marazita, M.L., Case/control family study of autonomic nervous system dysfunction in idiopathic congenital central hypoventilation syndrome, Am. J. Med. Genet. 100, 237–245, 2001. Schla¨fke, M.E., Kille, J.F. and Loeschcke, H.H., Elimination of central chemosensitivity by coagulation of a bilateral area on the ventral medullary surface in awake cats, Pflu¨ger Arch. 378, 231–241, 1979. Schla¨fke, M.E., See, W.R., Herker-See, A. and Loeschke, H.H., Respiratory response to hypoxia and hypercapnia after elimination of central chemosensitivity, Pflu¨ger Arch. 381, 241–248, 1979. Schafer, D. and Schla¨fke, M.E., Cardiorespiratory regulation in a model for Ondine’s curse syndrome, in Ventral Brainstem Mechanisms and Control of Respiration and Blood Pressure. Lung Biology in Health and Disease, Trouth, O.C., Millis, R.M., Kiwull-Scho¨ne, H.F. and Schla¨fke, M.E., eds., New York, Marcel Dekker, Inc., pp. 675–685, 1995. Burton, M.D., Kawashima, A., Brayer, J.A., Kazemi, H., Shannon, D.C., Schuchardt, A., Costantini, F., Pachnis, V. and Kinane, T.B., RET protooncogene is important for the development of respiratory CO2 sensitivity, J. Auton. Nerv. Syst. 63, 137–143, 1997. Aizenfisz, S., Dauger, S., Durand, E., Vardon, G., Levacher, B., Simonneau, M., Pachnis, V., Gaultier, C. and Gallego, J., Ventilatory responses to hypercapnia and hypoxia in heterozygous c-ret newborn mice, Respir. Physiol. Neurobiol. 131, 213–222, 2002. Erickson, J.T., Conover, J.C., Borday, V., Champagnat, J., Barbacid, M., Yancopoulos, G. and Katz, D.M., Mice lacking brain-derived neurotrophic factor exhibit visceral sensory neuron losses distinct from mice lacking NT4 and display a severe developmental deficit in control of breathing, J. Neurosci. 16, 5361–5371, 1996. Shirasawa, S., Arata, A., Onimaru, H., Roth, K.A., Brown, G.A., Horning, S., Arata, S., Okumura, K., Sasazuki, T. and Korsmeyer, S.J., Rnx deficiency results in congenital central hypoventilation, Nat. Genet. 24, 287–290, 2000. Amiel, J., Pelet, A., Trang, H., De Pontual, L., Simonneau, M., Munnich, A., Gaultier, C. and Lyonnet, S., Exclusion of RNX as a major gene in congenital central hypoventilation syndrome (CCHS, Ondine’s curse), Am. J. Med. Genet. 117A, 18–20, 2003.
310 77.
78.
79.
80.
81.
82. 83.
84.
85.
86.
87.
88.
89.
90.
Gozal Matera, I., Bachetti, T., Cinti, R., Lerone, M., Gagliardi, L., Morandi, F., Motta, M., Mosca, F., Ottonello, G., Piumelli, R., Schober, J.G., Ravazzolo, R. and Ceccherini, I., Mutational analysis of the RNX gene in congenital central hypoventilation syndrome, Am. J. Med. Genet. 113, 178–182, 2002. Folgering, H., Kuyper, F. and Kille, J.F., Primary alveolar hypoventilation (Ondine’s curse syndrome) in an infant without external arcuate nucleus. Case report, Bull. Eur. Physiopathol. Respir. 15, 659–665, 1979. Beal, M.F., Richardson, E.P., Jr., Brandstetter, R., Hedley-Whyte, E.T. and Hochberg, F.H., Localized brainstem ischemic damage and Ondine’s curse after near-drowning, Neurology 33, 717–721, 1983. Bogousslavsky, J., Khurana, R., Deruaz, J.P., Hornung, J.P., Regli, F., Janzer, R. and Perret, C., Respiratory failure and unilateral caudal brainstem infarction, Ann. Neurol. 28, 668–673, 1990. Liu, H.M., Loew, J.M. and Hunt, C.E., Congenital central hypoventilation syndrome: a pathologic study of the neuromuscular system, Neurology 28, 1013–1019, 1978. Jensen, T.H., Hansen, P.B. and Brodersen, P., Ondine’s curse in listeria monocytogenes brain stem encephalitis, Acta Neurol. Scand. 77, 505–506, 1988. Giangaspero, F., Schiavina, M., Sturani, C., Mondini, S. and Cirignotta, F., Failure of automatic control of ventilation (Ondine’s curse) associated with viral encephalitis of the brainstem: a clinicopathologic study of one case, Clin. Neuropathol. 7, 234–237, 1988. Weese-Mayer, D.E., Brouillette, R.T., Naidich, T.P., McClone, D.G. and Hunt, C.E., Magnetic resonance imaging and computerized tomography in central hypoventilation, Am. Rev. Respir. Dis. 137, 393–398, 1988. Cutz, E., Ma, T.K., Perrin, D.G., Moore, A.M. and Becker, L.E., Peripheral chemoreceptors in congenital central hypoventilation syndrome, Am. J. Respir. Crit. Care Med. 155, 358–363, 1997. Ogawa, S., Lee, T.M., Nayak, A.S. and Glynn, P., Oxygenation-sensitive contrast in magnetic resonance image of rodent brain at high magnetic fields, Magnet. Reson. Med. 14, 68–78, 1990. Hathout, G.M., Gambhir, S.S., Gopi, R.K., Kirlew, K.A.T., Choi, Y., So, G., Gozal, D., Harper, R.M., Lufkin, R.B. and Hawkins, R., A quantitative physiologic model of blood oxygenation for functional magnetic resonance imaging, Invest. Radiol. 30, 669–682, 1995. Gozal, D., Hathout, G.M., Kirlew, K.A.T., Tang, H., Woo, M.S., Zhang, J., Lufkin, R.B. and Harper, R.M., Localization of putative neural respiratory regions in the human by functional magnetic resonance imaging, J. Appl. Physiol. 76, 2076–2083, 1994. Gozal, D., Omidvar, O., Kirlew, K.A.T., Hathout, G.M., Hamilton, R., Lufkin, R.B. and Harper, R.M., Identification of human brain regions underlying responses to inspiratory loading with functional magnetic resonance imaging, Proc. Natl. Acad. Sci. USA 92, 6607–6611, 1995. Gozal, D., Omidvar, O., Kirlew, K.A.T., Hathout, G.M., Lufkin, R.B. and Harper, R.M., Brain regions mediating the response to resistive expiratory loads in humans, J. Clin. Invest. 97, 47–53, 1996.
Congenital Central Hypoventilation Syndrome 91. 92.
93.
94.
95.
96.
97.
98.
99.
100.
101.
102.
103.
104.
311
Gozal, D., Novel functional imaging strategies in assessment of respiratory control, Pediatr. Pulmonol. 23, 148–150, 1997. Spriggs, D., Saeed, M.M., Alger, J.R., Woo, M.A., Woo, M.S., Keens, T.G. and Harper, R.M., Neural responses to hypoxia in congenital central hypoventilation syndrome (CCHS) visualized by functional magnetic resonance imaging, Am. J. Resp. Crit. Care Med. 159, A587, 1999. Harper, R.M., Spriggs, D., Saeed, M.M., Alger, J.R., Woo, M.A., Woo, M.S., Gozal, D. and Keens, T.G., Functional magnetic resonance imaging during hypoxia challenges in congenital central hypoventilation syndrome (CCHS) reveals lateralized neural responses, Abstr. Soc. Neurosci. 25, A280, 1999. Spriggs, D., Saeed, M.M., Alger, J.R., Woo, M.A., Woo, M.S., Gozal, D., Keens, T.G. and Harper, R.M., Time course of functional magnetic resonance signal changes in response to hypercapnia in congenital central hypoventilation syndrome (CCHS), Abstr. Soc. Neurosci. 25, A280, 1999. Macey, K.E., Kuo, L., Kim, A., Yu, P.L., Woo, M.A., Saeed, M.M., Gozal, D. and Harper, R.M., Breathing patterns following hypercapnia in congenital central hypoventilation syndrome, Sleep 23, A16, 2000. Harper, R.M,. Yu, P.L., Saeed, M.M., Alger, J.R., Woo, M.A., Gozal, D. and Keens, T.G., Time trends of cerebellar fastigial nucleus responses to hypercapnia in congenital central hypoventilation syndrome (CCHS), Abstr. Soc. Neurosci. 26, A557, 2000. Paton, J.Y., Swaminathan, S., Sargent, C.W. and Keens, T.G., Hypoxic and hypercapneic ventilatory responses in awake children with congenital central hypoventilation syndrome, Am. Rev. Respir. Dis. 140, 368–372, 1989. Shea, S.A., Andres, L.P., Paydarfar, A., Banzett, R.B. and Shannon, D.C., Effect of mental activity on breathing in congenital central hypoventilation syndrome, Respir. Physiol. 94, 251–263, 1993. Gaultier, C., Trang-Pham, H., Praud, J.P. and Gallego, J., Cardiorespiratory control during sleep in the congenital central hypoventilation syndrome, Pediatr. Pulmonol. 23, 140–142, 1997. Gozal, D., Marcus, C.L., Shoseyov, D. and Keens, T.G., Peripheral chemoreceptor function in children with congenital central hypoventilation syndrome, J. Appl. Physiol. 74, 379–387, 1993. Shea, S.A., Andres, L.P., Shannon, D.C. and Banzett, R.B., Ventilatory responses to exercise in humans lacking ventilatory chemosensitivity, J. Physiol. (Lond.) 468, 623–640, 1993. Marcus, C.L., Bautista, D.B., Amihyia, A., Ward, S.L. and Keens, T.G., Hypercapneic arousal responses in children with congenital central hypoventilation syndrome, Pediatrics 88, 993–998, 1991. Silvestri, J.M., Weese-Mayer, D.E. and Flanagan, E.A., Congenital central hypoventilation syndrome: cardiorespiratory responses to moderate exercise, simulating daily activity, Pediatr. Pulmonol. 20, 89–93, 1995. Paton, J.Y., Swaminathan, S., Sargent, C.W., Hawksworth, A. and Keens, T.G., Ventilatory response to exercise in children with congenital central hypoventilation syndrome, Am. Rev. Resp. Dis. 147, 1185–1191, 1993.
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Gozal
105. Shea, S.A., Andres, L.P., Shannon, D.C. and Banzett, R.B., Ventilatory responses to exercise in humans lacking ventilatory chemosensitivity, J. Appl. Physiol. 68, 623–640, 1993. 106. Gozal, D., Marcus, C.L., Ward, S.L. and Keens, T.G., Ventilatory responses to passive leg motion in children with congenital central hypoventilation syndrome, Am. J. Resp. Crit. Care Med. 153, 761–768, 1996. 107. Gozal, D. and Simakajornboon, N., Passive motion of the extremities modifies alveolar ventilation during sleep in patients with CCHS, Am. J. Resp. Crit. Care Med. 162, 1747–1751, 2000. 108. Spengler, C.M., Gozal, D. and Shea, S.A., Chemoreceptive mechanisms elucidated by studies of congenital central hypoventilation syndrome, Respir. Physiol. 129, 247–255, 2001. 109. Hedner, J., Hedner, T., Breese, G.R., Lundell, K.H., Lundberg, D., Lundstro, N.R., Ostergaar, E., McCown, T.J. and Mueller, R.A., Changes in cerebrospinal fluid homovanillic acid in children with Ondine’s curse, Pediatr. Pulmonol. 3, 131–135, 1987. 110. Marazita, M.L., Maher, B.S., Cooper, M.E., Silvestri, J.M., Huffman, A.D., Smok-Pearsall, S.M., Kowal, M.H. and Weese-Mayer, D.E., Genetic segregation analysis of autonomic nervous system dysfunction in families of probands with idiopathic congenital central hypoventilation syndrome, Am. J. Med. Genet. 100, 229–236, 2001.
10 Upper Airway Obstruction in Sleep Apnea
SUSHEEL P. PATIL, HARTMUT SCHNEIDER, PHILIP L. SMITH, and ALAN R. SCHWARTZ Johns Hopkins University Baltimore, Maryland
I.
Introduction
Obstructive sleep apnea is a common disorder linked to the increasing prevalence of obesity in Western society, leading to recurrent oxyhemoglobin desaturations and arousals from sleep. Obstructive sleep apnea is caused by episodes of upper airway obstruction during sleep, and has been associated with increased morbidity and mortality [1–4] stemming from neurocognitive [5–8], cardiovascular [9–12], and respiratory dysfunction [13–18]. Upper airway obstruction is largely related to an increased propensity of the upper airway to collapse during sleep through a loss of neuromuscular tone. Sedatives and anesthetic agents can mimic the effects of natural sleep, thus increasing the risk for upper airway obstruction by blunting neuromuscular reflex and arousal responses that restore upper airway patency. This chapter will focus on the clinical and epidemiologic risk factors for upper airway obstruction, the pathophysiology of upper airway obstruction, and its implications for monitoring and treatment.
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Epidemiologic and Clinical Risk Factors
Obstructive sleep apnea is a comparatively new disorder with initial medical reports documenting the presence of discrete apneic episodes recurring throughout the night first appearing in the early 1960s [19–21]. While initial clinical reports established a strong association between sleep apnea and morbid obesity in patients with the Pickwickian syndrome [22–24], it was subsequently demonstrated that sleep apnea also occurred quite commonly in mildly to moderately overweight individuals who had no alterations in daytime gas exchange or evidence for cor pulmonale [13,25]. In a landmark publication in 1978, Remmers et al. [26] documented that apneas were due to the development of pharyngeal obstruction and were terminated by arousal from sleep. This report ushered in an era of physiologic investigations that examined the mechanism of upper airway obstruction [27–30], epidemiologic studies that defined the prevalence and risk factors for this disorder [31–42], and numerous investigations that examined the neurocognitive [5–8], behavioral [43–45], neuroendocrine [46–48], metabolic [25,49], and cardiovascular consequences [10,50,51] of sleep apnea. Investigations into the pathogenesis of sleep apnea and its consequences have employed standardized metrics of sleep apnea severity based on counts of apneas (no airflow) and hypopneas (reduced airflow associated with arousals and/or oxyhemoglobin desaturations) per hour of sleep (apnea–hypopnea index, AHI), associated arousals and oxyhemoglobin desaturations. With further refinements in monitoring technology, definitions of disease severity were expanded to include subjects with intermediate degrees of upper airway obstruction during sleep, and variable degrees of microarousals and oxyhemoglobin desaturations [52–59]. As standards for assessing the severity of upper airway obstruction and resulting alterations in gas exchange and sleep architecture have evolved, investigators have recognized a progression in upper airway obstruction that is characterized by snoring, obstructive hypopneas, and obstructive apnea [60]. In recent years, investigators have documented a high prevalence of sleep apnea in the general population. Early studies using a combination of symptom surveys and limited physiologic recordings in normal healthy adult populations suggested a high prevalence of sleep apnea in Western society [38,61–66]. In a subsequent, more comprehensive epidemiologic survey of sleep apnea in the general population, Young and co-workers [31] employed polysomnography to identify those with sleep apnea. The estimated prevalence of sleep apnea syndrome (AHI 5 and daytime hypersomnolence) was 2% in women and 4% in men for the population at large. Even higher point prevalence of sleep apnea (AHI 5) was observed in habitual snorers (9% of women and 24% of men). Furthermore, in an older, multicenter community-based cohort (the Sleep Heart Health Study) enriched with subjects who snored [9], sleep apnea was found in more than half of the
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men and a third of the women, indicating that the prevalence of this disorder may be quite high in older strata of the general population. Several recognized clinical risk factors have been associated with a greatly increased prevalence of obstructive sleep apnea in the general population [48,67]. Among these, obesity is thought to be the major risk factor. Mild to moderate obesity has been associated with a markedly increased prevalence of sleep apnea [25,31–42,68] and extreme obesity has been linked to increases in disease severity [69–73]. Male gender also constitutes a particularly strong risk factor, and confers a two- to three-fold increased risk of sleep apnea in the population at large [67,74]. It has been postulated that this increased risk is related to differences in the distribution of adipose tissue in men [35,48], who exhibit a predominantly central pattern of adiposity around the trunk and abdominal viscera, when compared with peripheral adiposity in women [75,76]. Increases in central adiposity with age may also account for an increase in sleep apnea prevalence in postmenopausal women [77–80]. In addition, recent studies have demonstrated familial aggregation and a racial predisposition to sleep apnea in individuals of African-American and Asian descent, suggesting that heritable factors also contribute to the development of sleep apnea [81–83]. Thus, obesity, and in particular central adiposity, may interact with male gender, age and genetic factors, leading to increases in the prevalence of sleep apnea. Nevertheless, the physiologic mechanisms linking obesity, male gender and sleep apnea have not been fully explained. Although age has also been considered a risk factor for sleep apnea, its effect may not be very pronounced. In a cross-sectional study of moderately obese, healthy male community subjects, no difference in the prevalence of sleep apnea was demonstrated across age groups [84]. Additional longitudinal data suggests that age does not contribute significantly to the prevalence of sleep apnea since there is minimal progression of disease in elderly cohorts over time [85]. Nevertheless, the high prevalence of sleep apnea in older populations may be related to age-related increases in body weight [86–89]. The pivotal role of obesity in the pathogenesis of sleep apnea is further suggested by the observation that modest decreases in body weight have been associated with substantial reductions in AHI [90,91]. Thus, both cross-sectional and longitudinal data suggest that obesity rather than age per se is a major risk factor for this disorder. With progressive increases in weight, patients develop signs of upper airway obstruction, including snoring, snorting, gasping, choking, and witnessed apneic episodes. As these symptoms progress, sleep disruption and daytime hypersomnolence typically ensue. When patients have been systematically queried in sleep disorders centers [92–98], investigators found that symptoms of upper airway obstruction, sleep disruption, and daytime hypersomnolence were highly predictive of the presence of sleep apnea. Of note, signs of upper airway obstruction were particularly predictive of
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this disorder in clinic-based populations. Moreover, when combined with measures of adiposity (i.e., body mass index or neck circumference) and male gender, symptoms of upper airway obstruction, sleep disruption, and excessive daytime somnolence were highly predictive of the presence of sleep apnea [96,97]. In addition to clinical and epidemiologic risk factors, obstructive sleep apnea has been associated with alterations in upper airway anatomy. Structural changes, including tonsillar hypertrophy [99–101], retrognathia [102–104], and variations in craniofacial structures [105–110], have been linked to an increased risk of sleep apnea, presumably by increasing upper airway collapsibility. Similarly, during wakefulness, computed tomographic and magnetic resonance imaging studies have demonstrated increased fatty tissue deposition and submucosal edema in the lateral walls of the pharynx, both of which narrow the pharyngeal lumen and may predispose to obstruction during sleep [19,111–113]. Underlying medical illness may also predispose to alterations in upper airway anatomy that play a role in the pathogenesis of upper airway obstruction. Endocrinopathy including testosterone administration [114–117], hypothyroidism [118–120], and acromegaly [121–123] are recognized risk factors for obstructive sleep apnea, and may produce structural changes in the neck and upper airway muscles that further predispose to upper airway obstruction. Specifically, testosterone replacement has been found to increase neck size, which has been associated with an increase in airway collapsibility [114,115,117,124]. Macroglossia, a recognized feature of hypothyroidism, acromegaly, and amyloidosis, may narrow the pharyngeal lumen and overload dilator muscles that maintain upper airway patency. Finally, hypothyroidism and acromegaly may result in soft tissue swelling and a concomitant myopathy that could narrow the pharynx and further compromise dilator muscle function, respectively. In recent years, clinical and epidemiologic evidence has also suggested a link between sleep apnea and underlying cardiovascular disease. It is now recognized that cardiovascular risk factors including male gender, visceral adiposity, glucose intolerance, and hypertension are highly associated with sleep apnea [32,34,48,125–127]. While some of these risk factors may predispose to sleep apnea (gender, obesity, and central adiposity), it also appears that cardiovascular and metabolic dysfunction are likely a consequence of this disorder. Evidence to support this association include a nearly three-fold increased incidence of hypertension in individuals with sleep apnea over a four-year follow-up period, suggesting that sleep apnea could be an antecedent for daytime hypertension [51,128]. Furthermore, sleep apnea has been associated with an increased risk of hypertension, stroke, angina, myocardial infarction and congestive heart failure in the ongoing multi-centered Sleep Heart Health Study [9,10]. Finally, among men with stable heart failure due to systolic dysfunction, approximately
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two-thirds of patients had sleep apnea [129–131]. Thus, the prevalence of sleep apnea may be particularly high in patients with underlying cardiovascular disease, and likely predisposes to increased cardiovascular morbidity and mortality. The cardiovascular sequelae of obstructive sleep apnea may be related to acute and chronic cardiovascular stress and metabolic dysregulation. Acutely, obstructive apneic episodes in experimental animals and humans are associated with recurrent surges in blood pressure following each apnea cycle [128,132–135]. Acute increases in blood pressure have been related to progressive increases in sympathetic drive throughout the apnea [136,137] and arousals at the termination of apneic episodes [134,137–139]. Investigators have further demonstrated that sleep apnea abolished the usual nocturnal decline in blood pressure [132,140], and that nocturnal blood pressure can rise above daytime levels in sleep apneic patients [140]. Experiments in animals have provided further insight into the mechanism for blood pressure elevations accompanying apneic episodes. The acute hemodynamic sequelae of sleep apnea have been shown to relate to the severity of oxyhemoglobin desaturations during the apneic episodes [141,142]. Blood pressure surges following apneas, however, appear to be caused by arousals and central effects of hypoxemia, which trigger pronounced sympathetic neural discharge and peripheral vasoconstriction, and increase cardiac afterload [137,143–145]. Chronic effects of intermittent hypoxemia and associated sympathetic neural discharges have now been linked to glucose intolerance [25,48,49,146], endothelial dysfunction [147–150], and disturbances in peripheral microvascular control [151]. Thus, evidence from animals and humans indicates that obstructive sleep apnea imposes significant acute and chronic metabolic and cardiovascular dysfunction. III.
Pathogenesis of Upper Airway Obstruction
A. Pathophysiology of Airflow Obstruction
Initial reports investigating the mechanism for upper airway obstruction [26] clearly demonstrated that the pharynx was the site of obstruction. The role of pharyngeal obstruction was further emphasized by the fact that obstructive sleep apnea could be eliminated by treatments that relieved or bypassed the upper airway [152–158]. Concepts concerning the pathogenesis of upper airway obstruction in obstructive sleep apnea were based on early work in normal anesthetized and paralyzed subjects [159,160], demonstrating that mechanical factors related to head, neck, and jaw position predisposed to pharyngeal obstruction. From fluoroscopic studies [161,162], it was shown that the tongue prolapsed during periods of obstruction, emphasizing that bulk soft tissue displacement was required
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for obstruction to occur. In particular, the tongue was noted to play a primary role, since the genioglossus muscle, the primary tongue protrusor, prolapsed into the pharynx whenever neuromuscular tone waned [26]. Further studies examined whether the obstruction occurred primarily at the retroglossal or retropalatal segment. Specifically, upper airway manometric studies during sleep demonstrated that wide pressure gradients developed in either pharyngeal segment during obstructive episodes, suggesting that collapse could occur in either the velo- or oropharynx [163–165]. Additional computed tomographic [166] and endoscopic imaging studies [167] have since documented that collapse can occur simultaneously in both pharyngeal segments, and can vary between non-REM and REM sleep stages within a single individual. These studies have given rise to the concept that while localized anatomic factors, such as adenotonsillar hypertrophy, retrognathia, and alterations in upper airway bony and soft tissue structures may predispose to collapse of certain pharyngeal segments, a more generalized defect in upper airway structural or neuromuscular control may be responsible for collapse along the entire length of the pharyngeal airway during sleep. To elucidate the mechanism of upper airway obstruction in obstructive sleep apnea, several approaches have been adopted to model the factors involved in the pathogenesis of pharyngeal collapse. Initial efforts focused on the interplay between extraluminal upper airway muscles that dilate and negative intraluminal pressures generated by the diaphragm that collapse the pharynx. It was originally postulated that upper airway patency was determined by the balance of pressures between the intraluminal and extraluminal spaces [26]. As intraluminal suction pressures overcame the dilating forces around the pharyngeal lumen, the theory held that the pharynx would progressively collapse and ultimately occlude during sleep. Later studies, however, have minimized the role of intraluminal suction pressures in the pathogenesis of upper airway obstruction by demonstrating that upper airway occlusion could occur spontaneously, even when intraluminal pressures were positive [168,169]. These observations resolved a major question regarding the role of negative intraluminal pressures in the pathogenesis of obstructive sleep apnea, and confirmed that negative pressures were not required for airway occlusion to occur. Rather, the markedly negative intraluminal pressures generated by the diaphragm during periods of upper airway obstruction were the consequence rather than the cause of upper airway occlusion. To further elucidate the mechanism for upper airway obstruction, investigators have examined airflow dynamics during periods of obstruction, and found that pressure-flow relationships were identical to those previously described for other collapsible biologic conduits, i.e., the Starling resistor (Figure 10.1) [170–172]. In a series of studies in sleeping humans
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Figure 10.1 The upper airway can be represented as a mechanical analogue (Starling resistor model) consisting of a rigid tube with a collapsible segment. Upstream (nasal) and downstream (hypopharyngeal) segments have fixed diameters and defined resistances, Rus and Rds, respectively. Pressures in these segments are represented by Pus and Pds, respectively. The collapsible segment has no resistance, but is subject to the surrounding pressure, Pcrit. Collapse occurs only when the surrounding pressure exceeds the intraluminal pressure (adapted from Ref. 60).
both with [29,173] and without sleep apnea [30], investigators demonstrated that upper airway flow dynamics resemble those seen in a Starling resistor in two respects [174]. First, as the downstream (tracheal) pressure falls during inspiration, inspiratory airflow attains a maximal level (VImax) and becomes independent of further decreases in downstream pressure (Figure 10.2A). This phenomenon, known as inspiratory airflow limitation, is characterized by a plateauing of airflow and is often associated with collapse of the pharynx and audible snoring. Second, under conditions of inspiratory airflow limitation, maximal inspiratory airflow varies linearly with changes in upstream nasal pressure, and falls from normal tidal airflow levels (300–500 ml/s) at higher levels of nasal pressure to zero as nasal pressure is lowered below the critical level (critical pressure, Pcrit) (Figure 10.2B). In fact, the critical closing pressure (Pcrit) is a direct measure of the collapsibility of the pharynx, and is operationally defined by lowering the nasal pressure until inspiratory airflow ceases (the airway occludes). In further studies, the upstream nasal pressure has been manipulated systematically, and critical pressures were measured in groups of individuals manifesting varying degrees of upper airway obstruction during sleep (Figure 10.3) [29,59,60]. Critical pressures were markedly negative in normal individuals with evidence of airflow obstruction, whereas critical pressures were positive in apneic patients with complete upper airway occlusion. In patients with partial airflow obstruction during sleep (obstructive hypopnea, upper airway resistance syndrome, and asymptomatic snorers), critical pressures were between these two extremes (minimally to moderately
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(A)
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Figure 10.2 (A) Initially as downstream pressure (Pesophageal) falls, inspiratory airflow increases. Inspiratory airflow reaches a maximal level (VImax) that is not exceeded as the downstream pressure continues to fall. As airflow plateaus (see arrows), inspiratory airflow limitation ensues and is characterized by airflow that is independent of further decreases in downstream pressure. (B) During conditions of inspiratory flow limitation, VImax directly varies with upstream nasal pressure (PN) and falls from normal tidal values to zero as PN is lowered below Pcrit (adapted from Ref. 174).
negative) [29,30,60]. These observations suggested that varying degrees of upper airway obstruction during sleep were associated with quantitative differences in critical pressures, reflecting differences in upper airway collapsibility across the spectrum from health to disease. Intervention studies have demonstrated that improvements in sleep apnea severity (AHI) are predicted by reductions in critical pressure after specific interventions. For example, in a study of weight loss intervention [69], decreases in critical pressure were directly correlated with decreases in weight, and sleep apnea remitted when critical pressures fell below 4 cmH2O. In a subsequent study examining the effect of uvulopalatopharyngoplasty on critical pressures and sleep apnea severity, it was shown that sleep apnea severity also correlated with reductions in critical pressure below a similar threshold of 4 cmH2O [175]. Finally, postural maneuvers, which lower the critical pressure by only 2–4 cmH2O, were associated with only modest improvements in sleep apnea severity for those in whom the critical pressure fell below atmospheric levels [176–178]. These intervention studies have clearly established that sleep apnea is due to elevations in critical pressure and that specific interventions can lead to predictable falls in
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Figure 10.3 Critical closing pressures of the upper airway (Pcrit) during sleep are plotted for groups of individuals that represent the clinical spectrum of obstructive sleep apnea—non-snoring, snoring, upper airway resistance syndrome (UARS), obstructive hypopnea, and obstructive apnea. Pcrit increases with increasing disease severity over a relatively narrow range of pressures. Note overlap between UARS and obstructive hypopneas, suggesting that the two disorders are indistinguishable in the degree of upper airway function, or in the impact of upper airway obstruction on sleep continuity (adapted from Refs. 30, 59 and 60).
critical pressures and improvements in sleep apnea that are based on the post-treatment value. Experimental studies have demonstrated that the critical pressure also determines the level of maximal inspiratory airflow. When the critical pressure is positive, airflow is zero at atmospheric nasal pressure. Progressive increases in nasal pressure in apneics have been associated with increasing levels of maximal inspiratory airflow and progression from obstructive apneas to hypopneas, to snoring and finally to normal breathing during sleep [29,60,173]. Once the nasal pressure exceeds the critical
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pressure, the airflow response to elevations in nasal pressure is determined by the airway resistance upstream to the site of pharyngeal collapse [173, 179]. Conversely, lowering the nasal pressure in normal individuals induces upper airway obstruction (inspiratory flow limitation) and progressive reductions in maximal inspiratory airflow [30]. As nasal pressure approaches a negative critical pressure in normal individuals, airflow falls and recurrent obstructive hypopneas and apneas ensue, leading to oxyhemoglobin desaturation and arousals from sleep [180]. In fact, a gradient of at least 5–8 cmH2O between the nasal and critical pressure is required to maintain tidal airflow levels, and approximately 10 cmH2O is required to abolish inspiratory airflow limitation completely [30,174,180]. These findings indicate that normal individuals and sleep apnea patients differ solely in their level of critical pressure, suggesting that obstructive sleep apnea is due to differences in upper airway collapsibility during sleep [180].
B. Anatomic Factors
While the critical pressure determines the degree of upper airway obstruction during sleep, the mechanisms leading to alterations in critical pressures are not well understood. In general, elevations in critical pressures have been attributed to either alterations in upper airway anatomical structures or disturbances in upper airway neuromuscular control [29,69]. Several approaches have been exploited to distinguish experimentally between these mechanisms for elevations in critical pressure. By eliminating neuromuscular activity with the administration of neuromuscular blocking agents, Isono and colleagues have recently examined the structural basis for alterations in critical pressure and compared values in normal individuals to those in patients with sleep apnea. In seminal studies [167,181–184], these investigators found elevated critical pressures in sleep apnea patients compared with normal controls, suggesting that alterations in upper airway anatomy contribute to the pathogenesis of upper airway obstruction. These investigators further determined that obesity, jaw position, acromegaly, tonsillar hypertrophy, and a smaller bony enclosure surrounding the pharynx may also elevate critical pressure [106,185–189]. Based on their observations (Figure 10.4), it has been proposed that crowding of soft tissue surrounding the pharynx could elevate the critical pressure by increasing the peri-pharyngeal pressure [106]. In addition to soft tissue crowding the pharyngeal lumen, cervical structures may exert axial forces that also influence upper airway collapsibility. In studies of airflow dynamics in the isolated upper airway of animals [190,191], investigators found that critical pressures fell
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markedly when the trachea was pulled caudally. This decrease was attributed to stretching and unfolding of redundant pharyngeal mucosa. Moreover, it was hypothesized that increases in tension within the pharyngeal wall mucosa would make it better able to withstand the compressive radial forces exerted by surrounding tissues. In further studies examining the interaction between radial and axial forces, Rowley et al. [191] demonstrated that axial traction amplifies the response of radial forces that either dilate or compress the airway (Figure 10.5). Additional evidence in animal and human studies indicate that axial tension increases during inspiration when the diaphragm moves caudally along with mediastinal and tracheal structures [192,193]. As the airway elongates, increases in axial tension have been associated with improvements in upper airway patency during inspiration [194]. Conversely, obesity, which decreases lung volumes and elevates the diaphragm, may predispose to upper airway obstruction through the loss of axial forces on the pharyngeal segment. Thus, structural factors leading to a dynamic interplay between axial and radial forces likely elevate critical pressures in obese individuals and modulate critical pressures throughout the respiratory cycle.
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Figure 10.5 (A) Tracheal and tongue displacement result in axial and radial forces that stabilize the upper airway. (B) Tongue displacement results in dilating forces that prevent upper airway collapse through decreases in critical pressure (Pcrit). (C) Tracheal displacement results in axial tension that can further stabilize the upper airway. (D) Axial traction can potentiate the effect of radial forces, thereby causing further decreases in Pcrit (Data from Ref. 191).
Several lines of evidence suggest obesity is associated with anatomic alterations that may also predispose to upper airway obstruction. In an isolated rabbit upper airway model, Koenig and Thach [195] found that applying lard-filled bags to the cervical area increased the critical pressure, suggesting that local deposition of cervical and peri-pharyngeal fat increases critical pressures through direct pressure on the upper airway. Similarly, obesity has been linked with elevations in neck circumference and increased amounts of peri-pharyngeal fat [196,197], which could narrow and compress the upper airway. In fact, in sleep apnea patients, MRI studies [112,198] have demonstrated greater amounts of peri-pharyngeal fat, correlating with sleep apnea severity. Finally, histologic studies of resected uvular tissue revealed greater amounts of submucosal fatty tissue in patients with obstructive sleep apnea [199]. These findings suggest that the compressive effects of fatty tissue deposited around the pharynx increase upper airway collapsibility, and possibly offset the effects of dilator muscles that maintain airway patency. Anatomic alterations may also account for increases in upper airway collapsibility in obese compared with non-obese individuals [69]. Critical pressures do not decrease with anterior mandibular displacement in obese individuals, a maneuver known to restore upper airway patency in normal
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individuals [187]. The lack of response with anterior mandibular displacement may be attributed to the fact that obesity, and in particular central obesity, has been associated with reductions in lung volume [200], known to increase upper airway collapsibility both in humans and in animals [190–193,201,202]. Thus, obesity imposes mechanical loads on both the upper airway and respiratory system that contribute to upper airway narrowing, obstruction, and collapse during sleep. C. Neuromuscular Factors Neuromuscular Reflexes
In addition to alterations in upper airway anatomy, disturbances in neuromuscular control may also contribute to the pathogenesis of upper airway obstruction during sleep. As neuromuscular activity wanes at sleep onset, its role in the maintenance of upper airway patency may differ between normal individuals and sleep apnea patients. The effect of neuromuscular activity on the critical pressure can be discerned during natural sleep and general anesthesia—with or without complete neuromuscular blockade [167,203,204]. When neuromuscular activity was reduced or eliminated under general anesthesia, critical pressures correlated with the presence and severity of upper airway obstruction during sleep. As seen in Figure 10.6, critical pressures in normal individuals and sleep apnea patients were near atmospheric, at levels known to be associated with severe upper airway obstruction during sleep. Since much lower (more negative) critical pressures are required to maintain upper airway patency during sleep in normal individuals [30], the marked critical pressure elevation in normal individuals during anesthesia and neuromuscular blockade suggests that neuromuscular mechanisms exert a major influence in the normal maintenance of upper airway patency during sleep (Figure 10.7). In contrast, critical pressures in sleep apnea patients do not differ markedly between the intact sleeping and anesthetized/paralyzed states [29,30,60,167]. Whereas neuromuscular mechanisms appear to stabilize the upper airway in normal sleeping individuals, this latter finding in sleep apnea patients suggests that a disturbance in neuromuscular control leads to upper airway obstruction during sleep. This disturbance may be due to a loss of neural drive to the upper airway musculature or to a reduced work efficiency of pharyngeal muscles that maintain upper airway patency. Current evidence suggests that compensatory increases in neuromuscular mechanisms play a critical role in maintaining upper airway patency in both humans and animals. Mezzanotte et al. [205] found that waking genioglossal EMG was elevated in sleep apnea patients when compared with normal controls, and postulated that apneic patients compensate for a narrower pharyngeal airway with higher levels of waking neuromuscular activity. Similarly, Hendricks and colleagues
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Figure 10.6 The apnea–hypopnea index during REM sleep demonstrates positive correlation with measurements of critical closing pressure (Pcrit) during general anesthesia, a state during which neuromuscular responses are minimized (reprinted with permission from Ref. 203).
[206,207] have studied the effects of anatomic airway narrowing in a bulldog model of sleep apnea, and found evidence for increased neuromuscular activity during sleep that may lead to muscle fiber injury in the cervical muscles. Series and co-workers [208] biopsied upper airway muscles and noted increased glycolytic capacity and alterations in the length tension characteristics of the muscularis uvulae in apneic patients. These alterations in upper airway muscle properties are consistent with the notion that neuromuscular recruitment during wakefulness may be required to compensate for mechanical loads in snoring and sleep apnea patients. Of note, critical pressures during wakefulness varied directly with physiologic and metabolic disturbances of the muscularis uvulae, suggesting that upper airway muscles still did not fully compensate for mechanical loads (Figure 10.8). Thus, it appears that a complex interaction between airway muscles and anatomic structures is required to maintain upper airway patency during wakefulness, and that a critical failure of compensatory neuromuscular mechanisms occurs during sleep.
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Upper airway obstruction is known to trigger various neuromuscular reflexes that activate upper airway dilator muscles (Figure 10.9) [205,209]. In animal studies, pulmonary and upper airway mechanoreceptors and chemoreceptors act individually and in combination to modify upper airway dilator muscle activity [210–217]. While these findings attest to the importance of neural activation, the precise effect of neural activity on upper airway function has not been well characterized. Chemoreceptor and mechanoreceptor reflexes [213,218,219] play a major role in regulating upper airway neuromuscular control and collapsibility, as emphasized by studies in which hypercapnia markedly increased neuromuscular activity and decreased the critical pressure [219]. Thus, studies in animals suggest that neuromuscular mechanisms can play an important role in stabilizing the upper airway, and that the loss of these protective mechanisms may predispose to upper airway obstruction during sleep.
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Figure 10.8 Airflow obstruction stimulates neuromuscular mechanisms that compensate for mechanical loads in individuals with sleep apnea relative to individuals without sleep apnea. As shown, (A) increases in length tension of the muscularis uvulae are associated with increases in critical pressures (Pcrit). In addition, (B) increases in anaerobic enzyme activity are associated with increases in Pcrit measured during wakefulness, providing further evidence of compensation for mechanical loads on the upper airway (Data from Ref. 208).
Studies examining neural EMG activity of various muscles have documented the influence of chemoreceptor and mechanoreceptor reflex responses on upper airway neuromuscular control during wakefulness [220,221]. Nevertheless, the influence of chemoreceptors responses during sleep is unclear, since marked hypercapnia is generally not observed during apneic episodes and the obstruction often persists in the face of significant hypoxemia. Moreover, it is now thought that mechanoreceptors may not respond appropriately to the markedly negative intraluminal pressure generated during periods of upper airway obstruction [211,222–227]. Early studies demonstrated that topical anesthesia of the pharyngeal mucosa increased the number of obstructive apneas and hypopneas during sleep in normal subjects and loud snorers, and/or increased the duration of apneic episodes [228–231]. In addition, it appears that upper airway sensory
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Figure 10.9 Upper airway obstruction can trigger various neuromuscular responses, as illustrated, to prevent upper airway collapse (Data from Ref. 299).
pathways may be impaired, since temperature, two-point discrimination and vibratory thresholds are disrupted in sleep apnea patients compared with normal individuals [232–234]. Sensory receptor dysfunction could also attenuate the response of upper airway dilator muscles to the markedly negative airway pressures generated during periods of upper airway obstruction. Further evidence for sensorimotor dysfunction is provided by graded histopathologic and immunochemical alterations in the palatopharyngeus and muscularis uvulae in sleep apnea patients, relative to asymptomatic snorers and normal individuals [235–239]. Findings of muscle fiber type redistribution and injury (fascicular atrophy and grouped atrophy in muscle fibers), have suggested that myopathic as well as sensory dysfunction may further compromise neuromuscular responses to upper airway obstruction. In recent studies involving tracheostomized patients [169,240], marked decreases in both tonic and phasic genioglossal activity were noted when patients breathed through their tracheostomy compared with breathing through their nares. This finding suggested that cyclic changes in pharyngeal pressures drive phasic firing of the genioglossus muscle during inspiration. More recent work has clearly demonstrated that genioglossal activity correlates with changes in pharyngeal pressure under a variety of experimental conditions, including one in which an iron lung was used to dissociate effects of tidal swings in intraluminal airway pressure from the
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intrinsic respiratory pattern generators. Akahoshi and colleagues [241] demonstrated strong correlations between genioglossal EMG activity and pharyngeal pressure, regardless of whether pressure swings were modulated by hypercapnia or passive breathing in the iron lung (Figure 10.10). These findings confirm that pressure-sensing mechanisms play a prominent role in modulating upper airway neuromuscular activity during wakefulness, although their role during sleep has not been well studied. In addition to changes in neuromuscular activity, a recent study [169] has established marked phasic modulation of upper airway collapsibility throughout the respiratory cycle. In sleeping tracheostomized apneic patients, markedly diminished (more negative) critical pressures were observed when patients breathed through their pharynx as opposed to their tracheostomy, suggesting that neuromuscular responses to negative airway pressure decreased collapsibility (Figure 10.11). In contrast, when patients breathed through their tracheostomies, the critical pressure increased and rose even further during expiration compared with inspiration. These observations in tracheostomized subjects suggest that phasic mechanisms stabilize upper airway patency and that phasic changes in airflow regimen, pressure, and genioglossus EMG activity play a role in dynamically modulating upper airway function during sleep.
Upper Airway Muscles
Which upper airway muscles might be responsible for maintaining upper airway patency? Initially, investigators focused on the role of the genioglossus, a prominent airway dilator muscle. The effect of the genioglossus muscle on upper airway stability has been examined by stimulating the genioglossus electrically in the isolated feline and canine upper airways. In these animal studies [218,242–247], electrical stimulation led to a significant fall in pharyngeal compliance and critical pressures, which stiffened and lowered airway collapsibility. Later studies by Fregosi and co-workers [248] confirmed that electrical stimulation of the genioglossus stabilized the airway, and that co-stimulating the tongue’s protrusor and retrusor muscles augmented this effect. In addition, it appears that both muscle groups play a significant role physiologically in stabilizing and maintaining airway patency, since these muscle groups are recruited simultaneously under hypercapnic and hypoxic conditions [249,250]. The observations in animals have been extended to studies in humans, demonstrating that stimulation of the lingual protrusor (genioglossus) selectively or in combination with retractor muscles reduces airflow obstruction during sleep [251–254]. Studies by Oliven and co-workers confirmed these findings and demonstrated improvements in upper airway flow dynamics in both animals and humans [244,255–259]
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Figure 10.11 Phasic modulation of upper airway collapsibility (Pcrit) is present throughout the respiratory cycle in sleeping patients with tracheostomy (n ¼ 6). Pcrit decreases during inspiration versus peak expiration during tracheostomy (solid bars) breathing, and falls even lower with nasal breathing. Effect of nasal (open bar) vs. tracheostomy breathing and respiratory phase (tracheostomy breathing: peak expiratory, end expiratory, and inspiratory) on Pcrit is illustrated. Values are means SE (from Data Ref. 169).
during electrical stimulation at various lingual sites. Nevertheless, electrical stimulation of the lingual musculature only partially relieved upper airway obstruction, and did not eliminate obstructive apneic episodes completely [253,254], suggesting that other muscles must also be recruited to restore upper airway patency and normalize tidal airflow. It is well known that the intrinsic muscles of the pharyngeal wall and the cervical strap muscles also play a role in the maintenance of airway patency. In the isolated feline upper airway model, studies by Kuna [260,261] and others [262,263] have demonstrated that pharyngeal constrictor muscles are recruited phasically in states of high ventilatory drive, and thus may reduce airway compliance (stiffen the airway). Moreover, responses in upper airway patency to constrictor muscle stimulation
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depend on pharyngeal luminal size. The pharyngeal constrictors were found to narrow the airway when widely patent, yet dilate when narrowed [264,265]. In further studies, specific oro- and velopharyngeal muscles appear to differentially control the patency of specific pharyngeal segments, suggesting that coordinated action of these muscles is required to stabilize the airway along its entire length. In addition to the genioglossus and intrinsic pharyngeal muscles, the cervical strap muscles also play a major role in supporting the airway, primarily by exerting caudal traction on the hyoid bone complex [266]. Thus, coordinated action of muscles that dilate, elongate and stiffen the pharyngeal wall may be required to optimally preserve airway patency.
IV.
Therapeutic Implications
Despite the complexity in delineating the structural and neuromuscular factors modulating upper airway patency, two basic approaches can be taken to relieve pharyngeal obstruction in patients with obstructive sleep apnea (Figure 10.12). Both are predicated on the notion that collapse and airflow obstruction are due to elevations in critical pressure. As previously discussed, continuous positive airway pressure (CPAP) has been designed to overcome the obstruction (positive critical pressure) by elevating upstream pressures at the nose or mouth [152,267]. CPAP has been a mainstay of therapy for obstructive sleep apnea for nearly 20 years, and is effective in relieving obstruction in patients with a broad range of critical pressures. An alternative therapeutic approach to therapy is to lower the critical pressure by augmenting the structural or neuromuscular mechanisms required for the maintenance of airway patency. Current approaches to correcting alterations in upper airway mechanics include weight loss [69,91,268,269], postural maneuvers [176–178], upper airway reconstructive surgery (uvulopalatopharyngoplasty, transpalatal resection, adenotonsillar resection) [101,270–272], and a variety of procedures designed to move the hyoid, mandible and maxillary bones anteriorly [273–276]. Of note, Isono and colleagues have shown that anterior mandibular displacement lowered the critical pressure significantly in lean normal individuals, an effect that is best explained by reductions in tissue pressure surrounding the airway. Nevertheless, anterior mandibular displacement had little effect in obese patients (Figure 10.13). Alternatively, compensatory neuromuscular mechanisms can be improved with maneuvers that increase the recruitment of muscles during sleep, thereby counteracting the sleep-dependent decline in neuromuscular activity leading to upper airway obstruction. Initial efforts to electrically stimulate specific upper airway muscles have offered partial relief of upper
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Figure 10.12 Upper airway obstruction may be relieved through either of two mechanisms. (A) Increases in nasal pressure (PN) beyond the critical pressure (Pcrit) of the upper airway will result in airflow. (B) Alternatively, reductions in tissue pressure through surgical intervention (e.g., UPPP [uvulopalatopharyngoplasty], LAUP [laser-assisted uvulopalatoplasty], mandibular advancement, etc.) or neuromuscular stimulation (e.g., hypoglossal nerve stimulation) will reduce Pcrit, resulting in a patent upper airway (Data from Ref. 300).
airway obstruction [251–254,277], as have pharmacologic strategies that stimulate upper airway motor neuron pools with tricyclic antidepressants [278–282] and serotonergic agents [283–290]. Refinements in these therapeutic strategies will undoubtedly require more precise elucidation of the neurochemical, sensorimotor, and mechanical factors that maintain upper airway patency during sleep. Therapeutic monitoring and intervention is particularly important in the peri-operative setting, where lingering effects of pharmacologic agents can depress upper airway neuromuscular activity and predispose to upper airway obstruction and obstructive sleep apnea. In general, monitoring should be instituted to detect airway obstruction arising from CNS depressants such as alcohol, benzodiazepines, opioids, and general anesthetics, which block compensatory neural responses in several ways. First, alcohol ingestion has been shown to decrease genioglossal muscle activity [291–293], thereby blunting reflex recruitment of the upper airway musculature and predisposing to worsening obstruction. Second, arousal
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responses to airway occlusion are known to be prolonged in normal sleeping subjects after alcohol ingestion [294]. Finally, the combined effects of CNS depressants on the upper airway musculature and arousal responses could account for observed increases in the frequency and duration of apneic episodes, and worsening of oxyhemoglobin desaturations after alcohol ingestion [295]. Of special note, sedation with midazolam has been associated with upper airway critical pressures equivalent to those during sleep in normal individuals [296,297]. In contrast, general anesthesia (isoflurane) has been associated with markedly elevated critical pressures approximating those found during complete neuromuscular blockade [167,203,298], suggesting that general anesthesia is associated with a nearly complete loss of protective upper airway reflex mechanisms. To avert complications arising from upper airway obstruction postoperatively, oximetry should be monitored, postural maneuvers (semi-recumbent posture) and nasal CPAP should be implemented, and doses of CNS depressants should be minimized.
V.
Summary and Conclusions
In conclusion, obstructive sleep apnea is a common disorder of Western society, particularly in overweight men and postmenopausal women.
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By modeling the upper airway as a simple collapsible conduit, investigators have determined that the severity of upper airway obstruction during sleep is related to quantitative differences in critical pressure, which reflect differences in the degree of upper airway collapsibility. Elevations in critical pressure appear to be due to a complex interaction of mechanical alterations and disturbances in upper airway neuromuscular control. In turn, disturbances in neuromuscular control and responses to upper airway obstruction lead to alterations in radial and axial traction on the pharyngeal airway, as well as variations in the neuromotor tone of the intrinsic muscles of specific pharyngeal segments. Understanding these pathophysiologic mechanisms can guide our approach to the treatment of obstructive sleep apnea. In the perioperative setting, awareness of the effects of anesthetic agents on the upper airway, the acute hemodynamic stresses produced by obstructive apneic episodes, and their associated desaturations and arousals make it particularly important to prevent and relieve airway obstruction when it occurs. References 1.
2.
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5.
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7.
8.
He, J., Kryger, M.H., Zorick, F.J., Conway, W. and Roth, T., Mortality and apnea index in obstructive sleep apnea. Experience in 385 male patients, Chest 94, 9–14, 1988. Lavie, P., Herer, P., Peled, R., Berger, I., Yoffe, N., Zomer, J. and Rubin, A.H., Mortality in sleep apnea patients: A multivariate analysis of risk factors, Sleep 18, 149–157, 1995. Bliwise, D.L., Bliwise, N.G., Partinen, M., Pursley, A.M. and Dement, W.C., Sleep apnea and mortality in an aged cohort, Am. J. Public Health 78, 544–547, 1988. Partinen, M., Jamieson, A. and Guilleminault, C., Long-term outcome for obstructive sleep apnea syndrome patients. Mortality, Chest 94, 1200–1204, 1988. Punjabi, N.M., O’Hearn, D.J., Neubauer, D.N., Nieto, F.J., Schwartz, A.R., Smith, P.L. and Bandeen-Roche, K., Modeling hypersomnolence in sleepdisordered breathing. A novel approach using survival analysis, Am. J. Respir. Crit. Care Med. 159, 1703–1709, 1999. Guilleminault, C., Partinen, M., Quera-Salva, M.A., Hayes, B., Dement, W.C. and Nino-Murcia, G., Determinants of daytime sleepiness in obstructive sleep apnea, Chest 94, 32–37, 1988. Roehrs, T., Zorick, F., Wittig, R., Conway, W. and Roth, T., Predictors of objective level of daytime sleepiness in patients with sleep-related breathing disorders, Chest 95, 1202–1206, 1989. Colt, H.G., Haas, H. and Rich, G.B., Hypoxemia vs. sleep fragmentation as cause of excessive daytime sleepiness in obstructive sleep apnea, Chest 100, 1542–1548, 1991.
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16. 17. 18.
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23.
337
Nieto, F.J., Young, T.B., Lind, B.K., Shahar, E., Samet, J.M., Redline, S., D’Agostino, R.B., Newman, A.B., Lebowitz, M.D. and Pickering, T.G., Association of sleep-disordered breathing, sleep apnea, and hypertension in a large community-based study. Sleep Heart Health Study, JAMA 283, 1829–1836, 2000. Shahar, E., Whitney, C.W., Redline, S., Lee, E.T., Newman, A.B., Javier, N.F., O’Connor, G.T., Boland, L.L., Schwartz, J.E. and Samet, J.M., Sleep-disordered breathing and cardiovascular disease: Cross-sectional results of the Sleep Heart Health Study, Am. J. Respir. Crit. Care Med. 163, 19–25, 2001. Newman, A.B., Nieto, F.J., Guidry, U., Lind, B.K., Redline, S., Pickering, T.G. and Quan, S.F., Relation of sleep-disordered breathing to cardiovascular disease risk factors: The Sleep Heart Health Study, Am. J. Epidemiol. 154, 50–59, 2001. Peker, Y., Hedner, J., Norum, J., Kraiczi, H. and Carlson, J., Increased incidence of cardiovascular disease in middle-aged men with obstructive sleep apnea: A 7-year follow-up, Am. J. Respir. Crit. Care Med. 166, 159–165, 2002. Gold, A.R., Schwartz, A.R., Wise, R.A. and Smith, P.L., Pulmonary function and respiratory chemosensitivity in moderately obese patients with sleep apnea, Chest 103, 1325–1329, 1993. Rapoport, D.M., Garay, S.M., Epstein, H. and Goldring, R.M., Hypercapnia in the obstructive sleep apnea syndrome. A reevaluation of the ‘Pickwickian syndrome’, Chest 89, 627–635, 1986. Bradley, T.D., Rutherford, R., Grossman, R.F., Lue, F., Zamel, N., Moldofsky, H. and Phillipson, E.A., Role of daytime hypoxemia in the pathogenesis of right heart failure in the obstructive sleep apnea syndrome, Am. Rev. Respir. Dis. 131, 835–839, 1985. Rajagopal, K.R., Abbrecht, P.H. and Tellis, C.J., Control of breathing in obstructive sleep apnea, Chest 85, 174–180, 1984. Bradley, T.D., Right and left ventricular functional impairment and sleep apnea. Clin. Chest Med. 13, 459–479, 1992. Krieger, J., Sforza, E., Apprill, M., Lampert, E., Weitzenblum, E. and Ratomaharo, J., Pulmonary hypertension, hypoxemia, and hypercapnia in obstructive sleep apnea patients, Chest 96, 729–737, 1989. Gastaut, H., Tassinari, C.A. and Duron, B., Polygraphic study of the episodic diurnal and nocturnal (hypnic and respiratory) manifestations of the Pickwick syndrome, Brain Res. 1, 167–186, 1966. Kuhlo, W., Doll, E. and Francj, M.D., Erfolgreiche behandlung eines Pickwick Syndroms durch eine dauertrachekanuele, Dtsch. Med. Wochenschr. 94, 1286–1290, 1964. Drachman, D.B. and Gumnit, R.J., Periodic alteration of consciousness in the ‘Pickwick’ Syndrome, Arch. Neurol. 6, 63–69, 1962. Burwell, C.S., Robin, E.D., Whaley, R.D. and Bickelmann, A.G., Extreme obesity associated with alveolar hypoventilation—Pickwickian syndrome, Am. J. Med. 21, 811–817, 1956. Addington, W.W., Pfeffer, S.H. and Gaensler, E.A., Obesity and alveolar hypoventilation, Respiration 26, 214–225, 1969.
338 24.
25.
26. 27. 28. 29.
30.
31.
32.
33.
34.
35.
36.
37.
38. 39.
Patil et al. Lugaresi, E., Coccagna, G., Mantovani, M., Cirignotta, F., Ambrosetto, G. and Baturic, P., Hypersomnia with periodic breathing: Periodic apneas and alveolar hypoventilation during sleep, Bull. Physiopathol. Respir. (Nancy) 8, 1103–1113, 1972. Punjabi, N.M., Sorkin, J.D., Katzel, L.I., Goldberg, A.P., Schwartz, A.R. and Smith, P.L., Sleep-disordered breathing and insulin resistance in middle-aged and overweight men, Am. J. Respir. Crit. Care Med. 165, 677–682, 2002. Remmers, J.E., deGroot, W.J., Sauerland, E.K. and Anch, A.M., Pathogenesis of upper airway occlusion during sleep, J. Appl. Physiol. 44, 931–938, 1978. Issa, F.G. and Sullivan, C.E., Upper airway closing pressures in snorers, J. Appl. Physiol. 57, 528–535, 1984. Issa, F.G. and Sullivan, C.E., Upper airway closing pressures in obstructive sleep apnea, J. Appl. Physiol. 57, 520–527, 1984. Smith, P.L., Wise, R.A., Gold, A.R., Schwartz, A.R. and Permutt, S., Upper airway pressure-flow relationships in obstructive sleep apnea, J. Appl. Physiol. 64, 789–795, 1988. Schwartz, A.R., Smith, P.L., Wise, R.A., Gold, A.R. and Permutt, S., Induction of upper airway occlusion in sleeping individuals with subatmospheric nasal pressure, J. Appl. Physiol. 64, 535–542, 1988. Young, T., Palta, M., Dempsey, J., Skatrud, J., Weber, S. and Badr, S., The occurrence of sleep-disordered breathing among middle-aged adults, N. Engl. J. Med. 328, 1230–1235, 1993. Grunstein, R., Wilcox, I., Yang, T.S., Gould, Y. and Hedner, J., Snoring and sleep apnoea in men: Association with central obesity and hypertension, Int. J. Obes. Relat. Metab. Disord. 17, 533–540, 1993. Davies, R.J., Ali, N.J. and Stradling, J.R., Neck circumference and other clinical features in the diagnosis of the obstructive sleep apnoea syndrome, Thorax 47, 101–105, 1992. Levinson, P.D., McGarvey, S.T., Carlisle, C.C., Eveloff, S.E., Herbert, P.N. and Millman, R.P., Adiposity and cardiovascular risk factors in men with obstructive sleep apnea, Chest 103, 1336–1342, 1993. Millman, R.P., Carlisle, C.C., McGarvey, S.T., Eveloff, S.E. and Levinson, P.D., Body fat distribution and sleep apnea severity in women, Chest 107, 362–366, 1995. Shinohara, E., Kihara, S., Yamashita, S., Yamane, M., Nishida, M., Arai, T., Kotani, K., Nakamura, T., Takemura, K. and Matsuzawa, Y., Visceral fat accumulation as an important risk factor for obstructive sleep apnoea syndrome in obese subjects, J. Intern. Med. 241, 11–18, 1997. Bearpark, H., Elliott, L., Grunstein, R., Hedner, J., Cullen, S., Schneider, H., Althaus, W. and Sullivan, C., Occurrence and correlates of sleep disordered breathing in the Australian town of Busselton: A preliminary analysis, Sleep 16, S3–S5, 1993. Stradling, J.R. and Crosby, J.H., Predictors and prevalence of obstructive sleep apnoea and snoring in 1001 middle aged men, Thorax 46, 85–90, 1991. Enright, P.L., Newman, A.B., Wahl, P.W., Manolio, T.A., Haponik, E.F. and Boyle, P.J., Prevalence and correlates of snoring and observed apneas in 5,201 older adults, Sleep 19, 531–538, 1996.
Upper Airway Obstruction 40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
51.
52.
339
Jennum, P. and Sjol, A., Epidemiology of snoring and obstructive sleep apnoea in a Danish population, age 30–60, J. Sleep Res. 1, 240–244, 1992. Olson, L.G., King, M.T., Hensley, M.J. and Saunders, N.A., A community study of snoring and sleep-disordered breathing. Prevalence, Am. J. Respir. Crit. Care Med. 152, 711–716, 1995. Ferini-Strambi, L., Zucconi, M., Palazzi, S., Castronovo, V., Oldani, A., Della, M.G. and Smirne, S., Snoring and nocturnal oxygen desaturations in an Italian middle-aged male population. Epidemiologic study with an ambulatory device, Chest 105, 1759–1764, 1994. Baldwin, C.M., Griffith, K.A., Nieto, F.J., O’Connor, G.T., Walsleben, J.A. and Redline, S., The association of sleep-disordered breathing and sleep symptoms with quality of life in the Sleep Heart Health Study, Sleep 24, 96–105, 2001. Weaver, T.E., Laizner, A.M., Evans, L.K., Maislin, G., Chugh, D.K., Lyon, K., Smith, P.L., Schwartz, A.R., Redline, S., Pack, A.I. and Dinges, D.F., An instrument to measure functional status outcomes for disorders of excessive sleepiness, Sleep 20, 835–843, 1997. Grunstein, R.R., Stenlof, K., Hedner, J.A. and Sjostrom, L., Impact of self-reported sleep-breathing disturbances on psychosocial performance in the Swedish Obese Subjects (SOS) Study, Sleep 18, 635–643, 1995. Grunstein, R.R., Handelsman, D.J., Lawrence, S.J., Blackwell, C., Caterson, I.D. and Sullivan, C.E., Neuroendocrine dysfunction in sleep apnea: Reversal by continuous positive airways pressure therapy, J. Clin. Endocrinol. Metab. 68, 352–358, 1989. Chin, K., Shimizu, K., Nakamura, T., Narai, N., Masuzaki, H., Ogawa, Y., Mishima, M., Nakao, K. and Ohi, M., Changes in intra-abdominal visceral fat and serum leptin levels in patients with obstructive sleep apnea syndrome following nasal continuous positive airway pressure therapy, Circulation 100, 706–712, 1999. Vgontzas, A.N., Papanicolaou, D.A., Bixler, E.O., Hopper, K., Lotsikas, A., Lin, H.M., Kales, A. and Chrousos, G.P., Sleep apnea and daytime sleepiness and fatigue: Relation to visceral obesity, insulin resistance, and hypercytokinemia, J. Clin. Endocrinol. Metab. 85, 1151–1158, 2000. Ip, M.S., Lam, B., Ng, M.M., Lam, W.K., Tsang, K.W. and Lam, K.S., Obstructive sleep apnea is independently associated with insulin resistance, Am. J. Respir. Crit. Care Med. 165, 670–676, 2002. Young, T., Peppard, P., Palta, M., Hla, K.M., Finn, L., Morgan, B. and Skatrud, J., Population-based study of sleep-disordered breathing as a risk factor for hypertension, Arch. Intern. Med. 157, 1746–1752, 1997. Peppard, P.E., Young, T., Palta, M. and Skatrud, J., Prospective study of the association between sleep-disordered breathing and hypertension, N. Engl. J. Med. 342, 1378–1384, 2000. Tsai, W.H., Flemons, W.W., Whitelaw, W.A. and Remmers, J.E., A comparison of apnea–hypopnea indices derived from different definitions of hypopnea, Am. J. Respir. Crit. Care Med. 159, 43–48, 1999.
340 53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63. 64.
65.
66.
Patil et al. Meoli, A.L., Casey, K.R., Clark, R.W., Coleman, J.A., Jr., Fayle, R.W., Troell, R.J. and Iber, C., Hypopnea in sleep-disordered breathing in adults, Sleep 24, 469–470, 2001. Atlas Task Force, American Sleep Disorders Association, EEG arousals: Scoring rules and examples: A preliminary report from the Sleep Disorders Atlas Task Force of the American Sleep Disorders Association, Sleep 15, 173–184, 1992. American Academy of Sleep Medicine, Sleep-related breathing disorders in adults: Recommendations for syndrome definition and measurement techniques in clinical research. The Report of an American Academy of Sleep Medicine Task Force, Sleep 22, 667–689, 1999. Ayappa, I., Norman, R.G., Krieger, A.C., Rosen, A., O’malley, R.L. and Rapoport, D.M., Non-invasive detection of respiratory effort-related arousals (RERAs) by a nasal cannula/pressure transducer system, Sleep 23, 763–771, 2000. Hosselet, J.J., Norman, R.G., Ayappa, I. and Rapoport, D.M., Detection of flow limitation with a nasal cannula/pressure transducer system, Am. J. Respir. Crit. Care Med. 157, 1461–1467, 1998. Guilleminault, C., Stoohs, R., Clerk, A., Cetel, M. and Maistros, P., A cause of excessive daytime sleepiness. The upper airway resistance syndrome, Chest 104, 781–787, 1993. Gold, A.R., Marcus, C.L., Dipalo, F. and Gold, M.S., Upper airway collapsibility during sleep in upper airway resistance syndrome, Chest 121, 1531–1540, 2002. Gleadhill, I.C., Schwartz, A.R., Schubert, N., Wise, R.A., Permutt, S. and Smith, P.L., Upper airway collapsibility in snorers and in patients with obstructive hypopnea and apnea, Am. Rev. Respir. Dis. 143, 1300–1303, 1991. Lavie, P., Incidence of sleep apnea in a presumably healthy working population: A significant relationship with excessive daytime sleepiness, Sleep 6, 312–318, 1983. Franceschi, M., Zamproni, P., Crippa, D. and Smirne, S., Excessive daytime sleepiness: A 1-year study in an unselected inpatient population, Sleep 5, 239–247, 1982. Gislason, T. and Taube, A., Prevalence of sleep apnea syndrome—estimation by two stage sampling, Ups. J. Med. Sci. 92, 193–203, 1987. Cirignotta, F., D’Alessandro, R., Partinen, M., Zucconi, M., Cristina, E., Gerardi, R., Cacciatore, F.M. and Lugaresi, E., Prevalence of every night snoring and obstructive sleep apnoeas among 30–69-year-old men in Bologna, Italy, Acta Neurol. Scand. 79, 366–372, 1989. Schmidt-Nowara, W.W., Coultas, D.B., Wiggins, C., Skipper, B.E. and Samet, J.M., Snoring in a Hispanic-American population. Risk factors and association with hypertension and other morbidity, Arch. Intern. Med. 150, 597–601, 1990. Gislason, T., Benediktsdottir, B., Bjornsson, J.K., Kjartansson, G., Kjeld, M. and Kristbjarnarson, H., Snoring, hypertension, and the sleep apnea syndrome. An epidemiologic survey of middle-aged women, Chest 103, 1147– 1151, 1993.
Upper Airway Obstruction 67.
68.
69.
70.
71.
72.
73.
74. 75.
76. 77.
78.
79. 80.
81.
82.
341
Young, T., Peppard, P.E. and Gottlieb, D.J., Epidemiology of obstructive sleep apnea: A population health perspective, Am. J. Respir. Crit. Care Med. 165, 1217–1239, 2002. Peppard, P.E., Young, T., Palta, M., Dempsey, J. and Skatrud, J., Longitudinal study of moderate weight change and sleep-disordered breathing, JAMA 284, 3015–3021, 2000. Schwartz, A.R., Gold, A.R., Schubert, N., Stryzak, A., Wise, R.A., Permutt, S. and Smith, P.L., Effect of weight loss on upper airway collapsibility in obstructive sleep apnea, Am. Rev. Respir. Dis. 144, 494–498, 1991. Rajala, R., Partinen, M., Sane, T., Pelkonen, R., Huikuri, K. and Seppalainen, A.M., Obstructive sleep apnoea syndrome in morbidly obese patients, J. Intern. Med. 230, 125–129, 1991. Peiser, J., Lavie, P., Ovnat, A. and Charuzi, I., Sleep apnea syndrome in the morbidly obese as an indication for weight reduction surgery, Ann. Surg. 199, 112–115, 1984. Serafini, F.M., MacDowell, A.W., Rosemurgy, A.S., Strait, T. and Murr, M.M., Clinical predictors of sleep apnea in patients undergoing bariatric surgery, Obes. Surg. 11, 28–31, 2001. Resta, O., Foschino-Barbaro, M.P., Legari, G., Talamo, S., Bonfitto, P., Palumbo, A., Minenna, A., Giorgino, R. and De Pergola, G., Sleep-related breathing disorders, loud snoring and excessive daytime sleepiness in obese subjects, Int. J. Obes. Relat. Metab. Disord. 25, 669–675, 2001. Strohl, K.P. and Redline, S., Recognition of obstructive sleep apnea, Am. J. Resp. Crit. Care Med. 154, 279–289, 1996. Ledoux, M., Lambert, J., Reeder, B.A. and Despres, J.P., Correlation between cardiovascular disease risk factors and simple anthropometric measures. Canadian Heart Health Surveys Research Group, CMAJ 157(Suppl. 1), S46–S53, 1997. Legato, M.J., Gender-specific aspects of obesity, Int. J. Fertil. Womens Med. 42, 184–197, 1997. Guilleminault, C., Stoohs, R., Kim, Y.D., Chervin, R., Black, J. and Clerk, A., Upper airway sleep-disordered breathing in women, Ann. Intern. Med. 122, 493–501, 1995. Wilhoit, S.C. and Suratt, P.M., Obstructive sleep apnea in premenopausal women. A comparison with men and with postmenopausal women, Chest 91, 654–658, 1987. Guilleminault, C., Quera-Salva, M.A., Partinen, M. and Jamieson, A., Women and the obstructive sleep apnea syndrome, Chest 93, 104–109, 1988. Block, A.J., Wynne, J.W. and Boysen, P.G., Sleep-disordered breathing and nocturnal oxygen desaturation in postmenopausal women, Am. J. Med. 69, 75– 79, 1980. Redline, S., Tishler, P.V., Hans, M.G., Tosteson, T.D., Strohl, K.P. and Spry, K., Racial differences in sleep-disordered breathing in African-Americans and Caucasians, Am. J. Respir. Crit. Care Med. 155, 186–192, 1997. [Published erratum appears in Am. J. Respir. Crit. Care Med. 155, 1820, 1997]. Redline, S., Tishler, P.V., Schluchter, M., Aylor, J., Clark, K. and Graham, G., Risk factors for sleep-disordered breathing in children. Associations with
342
83.
84. 85.
86.
87.
88.
89.
90.
91.
92. 93.
94. 95.
96.
97. 98.
Patil et al. obesity, race, and respiratory problems, Am. J. Respir. Crit. Care Med. 159, 1527–1532, 1999. Redline, S., Tosteson, T., Tishler, P.V. and Carskadon, M.A., Studies in the genetics of obstructive sleep apnea. Familial aggregation of symptoms associated with sleep-related breathing disturbances, Am. Rev. Respir. Dis. 145, 440–444, 1992. Smith, P.L., Meyers, D.A. and Schubert, N.M., Assessment of sleep apnea severity in the elderly, Gerontologist 25, 119, 1985. Bliwise, D., Carskadon, M., Carey, E. and Dement, W., Longitudinal development of sleep-related respiratory disturbance in adult humans, J. Gerontol. 39, 290–293, 1984. Lahti-Koski, M., Vartiainen, E., Mannisto, S. and Pietinen, P., Age, education and occupation as determinants of trends in body mass index in Finland from 1982 to 1997, Int. J. Obes. Relat. Metab. Disord. 24, 1669–1676, 2000. Lahti-Koski, M., Pietinen, P., Mannisto, S. and Vartiainen, E., Trends in waist-to-hip ratio and its determinants in adults in Finland from 1987 to 1997, Am. J. Clin. Nutr. 72, 1436–1444, 2000. Tahara, Y., Moji, K., Aoyagi, K., Tsunawake, N., Muraki, S. and MascieTaylor, C.G., Age-related pattern of body density and body composition of Japanese men and women 18–59 years of age, Am. J. Hum. Biol. 14, 743–752, 2002. Welon, Z., Szklarska, A., Bielicki, T. and Malina, R.M., Sex differences in the pattern of age-dependent increase in the BMI from 20–59 years, Am. J. Hum. Biol. 14, 693–698, 2002. Browman, C.P., Sampson, M.G., Yolles, S.F., Gujavarty, K.S., Weiler, S.J., Walsleben, J.A., Hahn, P.M. and Mitler, M.M., Obstructive sleep apnea and body weight, Chest 85, 435–438, 1984. Smith, P.L., Gold, A.R., Meyers, D.A., Haponik, E.F. and Bleecker, E.R., Weight loss in mildly to moderately obese patients with obstructive sleep apnea, Ann. Intern. Med. 103, 850–855, 1985. Rowley, J.A., Aboussouan, L.S. and Badr, M.S., The use of clinical prediction formulas in the evaluation of obstructive sleep apnea, Sleep 23, 929–938, 2000. Netzer, N.C., Stoohs, R.A., Netzer, C.M., Clark, K. and Strohl, K.P., Using the Berlin Questionnaire to identify patients at risk for the sleep apnea syndrome, Ann. Intern. Med. 131, 485–491, 1999. Flemons, W.W. and Remmers, J.E., The diagnosis of sleep apnea: Questionnaires and home studies, Sleep 19, S243–S247, 1996. Flemons, W.W., Whitelaw, W.A., Brant, R. and Remmers, J.E., Likelihood ratios for a sleep apnea clinical prediction rule, Am. J. Respir. Crit. Care Med. 150, 1279–1285, 1994. Maislin, G., Pack, A.I., Kribbs, N.B., Smith, P.L., Schwartz, A.R., Kline, L.R., Schwab, R.J. and Dinges, D.F., A survey screen for prediction of apnea, Sleep 18, 158–166, 1995. Viner, S., Szalai, J.P. and Hoffstein, V., Are history and physical examination a good screening test for sleep apnea? Ann. Intern. Med. 115, 356–359, 1991. Crocker, B.D., Olson, L.G., Saunders, N.A., Hensley, M.J., McKeon, J.L., Allen, K.M. and Gyulay, S.G., Estimation of the probability of disturbed
Upper Airway Obstruction
99. 100.
101.
102.
103.
104.
105.
106.
107.
108.
109.
110.
111.
112.
343
breathing during sleep before a sleep study, Am. Rev. Respir. Dis. 142, 14–18, 1990. Moser, R.J., III and Rajagopal, K.R., Obstructive sleep apnea in adults with tonsillar hypertrophy, Arch. Intern. Med. 147, 1265–1267, 1987. Miyazaki, S., Itasaka, Y., Tada, H., Ishikawa, K. and Togawa, K., Effectiveness of tonsillectomy in adult sleep apnea syndrome, Psychiatry Clin. Neurosci. 52, 222–223, 1998. Verse, T., Kroker, B.A., Pirsig, W. and Brosch, S., Tonsillectomy as a treatment of obstructive sleep apnea in adults with tonsillar hypertrophy, Laryngoscope 110, 1556–1559, 2000. Riley, R., Guilleminault, C., Herran, J. and Powell, N., Cephalometric analyses and flow-volume loops in obstructive sleep apnea patients, Sleep 6, 303–311, 1983. Lyberg, T., Krogstad, O. and Djupesland, G., Cephalometric analysis in patients with obstructive sleep apnoea syndrome. I. Skeletal morphology, J. Laryngol. Otol. 103, 287–292, 1989. Tangugsorn, V., Skatvedt, O., Krogstad, O. and Lyberg, T., Obstructive sleep apnoea: A cephalometric study. Part I. Cervico-craniofacial skeletal morphology, Eur. J. Orthod. 17, 45–56, 1995. Millman, R.P., Acebo, C., Rosenberg, C. and Carskadon, M.A., Sleep, breathing, and cephalometrics in older children and young adults. Part II— Response to nasal occlusion, Chest 109, 673–679, 1996. Watanabe, T., Isono, S., Tanaka, A., Tanzawa, H. and Nishino, T. Contribution of body habitus and craniofacial characteristics to segmental closing pressures of the passive pharynx in patients with sleep-disordered breathing, Am. J. Respir. Crit. Care Med. 165, 260–265, 2002. Paoli, J.R., Lauwers, F., Lacassagne, L., Tiberge, M., Dodart, L. and Boutault, F., Craniofacial differences according to the body mass index of patients with obstructive sleep apnoea syndrome: Cephalometric study in 85 patients, Br. J. Oral Maxillofac. Surg. 39, 40–45, 2001. Cistulli, P.A., Gotsopoulos, H. and Sullivan, C.E., Relationship between craniofacial abnormalities and sleep-disordered breathing in Marfan’s syndrome, Chest 120, 1455–1460, 2001. Ito, D., Akashiba, T., Yamamoto, H., Kosaka, N. and Horie, T., Craniofacial abnormalities in Japanese patients with severe obstructive sleep apnoea syndrome, Respirology 6, 157–161, 2001. Cakirer, B., Hans, M.G., Graham, G., Aylor, J., Tishler, P.V. and Redline, S., The relationship between craniofacial morphology and obstructive sleep apnea in whites and in African-Americans, Am. J. Respir. Crit. Care Med. 163, 947–950, 2001. Haponik, E.F., Smith, P.L., Bohlman, M.E., Allen, R.P., Goldman, S.M. and Bleecker, E.R., Computerized tomography in obstructive sleep apnea. Correlation of airway size with physiology during sleep and wakefulness, Am. Rev. Respir. Dis. 127, 221–226, 1983. Schwab, R.J., Gupta, K.B., Gefter, W.B., Metzger, L.J., Hoffman, E.A. and Pack, A.I., Upper airway and soft tissue anatomy in normal subjects and patients with sleep-disordered breathing. Significance of
344
113.
114.
115.
116.
117.
118.
119.
120. 121. 122. 123.
124.
125.
126.
127.
Patil et al. the lateral pharyngeal walls, Am. J. Respir. Crit. Care Med. 152, 1673–1689, 1995. Trudo, F.J., Gefter, W.B., Welch, K.C., Gupta, K.B., Maislin, G. and Schwab, R.J., State-related changes in upper airway caliber and surrounding soft-tissue structures in normal subjects, Am. J. Respir. Crit. Care Med. 158, 1259–1270, 1998. Sandblom, R.E., Matsumoto, A.M., Schoene, R.B., Lee, K.A., Giblin, E.C., Bremner, W.J. and Pierson, D.J., Obstructive sleep apnea syndrome induced by testosterone administration, N. Engl. J. Med. 308, 508–510, 1983. Schneider, B.K., Pickett, C.K., Zwillich, C.W., Weil, J.V., McDermott, M.T., Santen, R.J., Varano, L.A. and White, D.P., Influence of testosterone on breathing during sleep, J. Appl. Physiol. 61, 618–623, 1986. Matsumoto, A.M., Sandblom, R.E., Schoene, R.B., Lee, K.A., Giblin, E.C., Pierson, D.J. and Bremner, W.J., Testosterone replacement in hypogonadal men: Effects on obstructive sleep apnoea, respiratory drives, and sleep, Clin. Endocrinol. (Oxf.) 22, 713–721, 1985. Cistulli, P.A., Grunstein, R.R. and Sullivan, C.E., Effect of testosterone administration on upper airway collapsibility during sleep, Am. J. Respir. Crit. Care Med. 149, 530–532, 1994. Rajagopal, K.R., Abbrecht, P.H., Derderian, S.S., Pickett, C., Hofeldt, F., Tellis, C.J. and Zwillich, C.W., Obstructive sleep apnea in hypothyroidism, Ann. Intern. Med. 101, 491–494, 1984. Kapur V.K., Koepsell T.D., deMaine, J., Hert, R., Sandblom, R.E. and Psaty, B.M., Association of hypothyroidism and obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 158, 1379–1383, 1998. Skjodt, N.M., Atkar, R. and Easton, P.A., Screening for hypothyroidism in sleep apnea, Am. J. Respir. Crit. Care Med. 160, 732–735, 1999. Mezon, B.J., West, P., MaClean, J.P. and Kryger, M.H., Sleep apnea in acromegaly, Am. J. Med. 69, 615–618, 1980. Grunstein, R.R., Ho, K.Y. and Sullivan, C.E., Sleep apnea in acromegaly, Ann. Intern. Med. 115, 527–532, 1991. Rosenow, F., Reuter, S., Deuss, U., Szelies, B., Hilgers, R.D., Winkelmann, W. and Heiss, W.D., Sleep apnoea in treated acromegaly: Relative frequency and predisposing factors, Clin. Endocrinol. (Oxf.) 45, 563–569, 1996. Johnson, M.W., Anch, A.M. and Remmers, J.E., Induction of the obstructive sleep apnea syndrome in a woman by exogenous androgen administration, Am. Rev. Respir. Dis. 129, 1023–1025, 1984. Bixler, E.O., Vgontzas, A.N., Lin, H.M., Ten Have, T., Leiby, B.E., Vela-Bueno, A. and Kales, A., Association of hypertension and sleepdisordered breathing, Arch. Intern. Med. 160, 2289–2295, 2000. Grunstein, R., Endocrine and metabolic disturbances in obstructive sleep apnea, in Sleep and Breathing Saunders, N.A. and Sullivan, C.E., eds, New York, Marcel Dekker, Inc., pp. 449–491, 1994. Wilcox, I., McNamara, S.G., Collins, F.L., Grunstein, R.R. and Sullivan, C.E., ‘Syndrome Z’: The interaction of sleep apnoea, vascular risk factors and heart disease, Thorax 53(Suppl. 3), S25–S28, 1998.
Upper Airway Obstruction
345
128. Davies, C.W., Crosby, J.H., Mullins, R.L., Barbour, C., Davies, R.J. and Stradling, J.R., Case-control study of 24 hour ambulatory blood pressure in patients with obstructive sleep apnoea and normal matched control subjects, Thorax 55, 736–740, 2000. 129. Javaheri, S., Parker, T.J., Liming, J.D., Corbett, W.S., Nishiyama, H., Wexler, L. and Roselle, G.A., Sleep apnea in 81 ambulatory male patients with stable heart failure. Types and their prevalence, consequences, and presentations, Circulation 97, 2154–2159, 1998. 130. Javaheri, S., Prevalence and prognostic significance of sleep apnea in heart failure, in Sleep Apnea: Implications in Cardiovascular and Cerebrovascular Disease, Lenfant, C., Bradly, T.D. and Floras, J.S., eds., New York, Marcel Dekker, Inc., pp. 415–433, 2000. 131. Sin, D.D., Fitzgerald, F., Parker, J.D., Newton, G., Floras, J.S. and Bradley, T.D., Risk factors for central and obstructive sleep apnea in 450 men and women with congestive heart failure, Am. J. Respir. Crit. Care Med. 160, 1101–1106, 1999. 132. Akashiba, T., Minemura, H., Yamamoto, H., Kosaka, N., Saito, O. and Horie, T., Nasal continuous positive airway pressure changes blood pressure ‘non-dippers’ to ‘dippers’ in patients with obstructive sleep apnea, Sleep 22, 849–853, 1999. 133. Davies, R.J., Vardi-Visy, K., Clarke, M. and Stradling, J.R., Identification of sleep disruption and sleep disordered breathing from the systolic blood pressure profile, Thorax 48, 1242–1247, 1993. 134. Davies, R.J., Belt, P.J., Roberts, S.J., Ali, N.J. and Stradling, J.R., Arterial blood pressure responses to graded transient arousal from sleep in normal humans, J. Appl. Physiol. 74, 1123–1130, 1993. 135. Wilcox, I., Grunstein, R.R., Hedner, J.A., Doyle, J., Collins, F.L., Fletcher, P.J., Kelly, D.T. and Sullivan, C.E., Effect of nasal continuous positive airway pressure during sleep on 24-hour blood pressure in obstructive sleep apnea, Sleep 16, 539–544, 1993. 136. Bao, G., Metreveli, N. and Fletcher, E.C., Acute and chronic blood pressure response to recurrent acoustic arousal in rats, Am. J. Hypertens. 12, 504–510, 1999. 137. O’Donnell, C.P., Ayuse, T., King, E.D., Schwartz, A.R., Smith, P.L. and Robotham, J.L., Airway obstruction during sleep increases blood pressure without arousal, J. Appl. Physiol. 80, 773–781, 1996. 138. O’Donnell, C.P., King, E.D., Schwartz, A.R., Robotham, J.L. and Smith, P.L., Relationship between blood pressure and airway obstruction during sleep in the dog, J. Appl. Physiol. 77, 1819–1828, 1994. 139. Schaub, C.D., Schneider, H. and O’Donnell, C.P., Mechanisms of acute and chronic blood pressure elevation in animal models of obstructive sleep apnea, in Sleep Apnea: Implications in Cardiovascular and Cerebrovascular Disease, Lenfant, C., Bradly, T.D. and Floras, J.S., eds., New York, Marcel Dekker, Inc., pp. 159–179, 2000. 140. Ali, N.J., Davies, R.J., Fleetham, J.A. and Stradling, J.R., The acute effects of continuous positive airway pressure and oxygen administration on blood pressure during obstructive sleep apnea, Chest 101, 1526–1532, 1992.
346
Patil et al.
141. Chen, L., Sica, A.L., Greenberg, H. and Scharf, S.M., Role of hypoxemia and hypercapnia in acute cardiovascular response to periodic apneas in sedated pigs, Respir. Physiol. 111, 257–269, 1998. 142. Chen, L., Sica, A.L. and Scharf, S.M., Mechanisms of acute cardiovascular response to periodic apneas in sedated pigs, J. Appl. Physiol. 86, 1236–1246, 1999. 143. Brooks, D., Horner, R.L., Kozar, L.F., Render-Teixeira, C.L. and Phillipson, E.A., Obstructive sleep apnea as a cause of systemic hypertension. Evidence from a canine model, J. Clin. Invest. 99, 106–109, 1997. 144. Horner, R.L., Brooks, D., Kozar, L.F., Tse, S. and Phillipson, E.A., Immediate effects of arousal from sleep on cardiac autonomic outflow in the absence of breathing in dogs, J. Appl. Physiol. 79, 151–162, 1995. 145. Schneider, H., Schaub, C.D., Chen, C.A., Andreoni, K.A., Schwartz, A.R., Smith, P.L., Robotham, J.L. and O’Donnell, C.P., Effects of arousal and sleep state on systemic and pulmonary hemodynamics in obstructive apnea, J. Appl. Physiol. 88, 1084–1092, 2000. 146. Stoohs, R.A., Facchini, F. and Guilleminault, C., Insulin resistance and sleep-disordered breathing in healthy humans, Am. J. Respir. Crit. Care Med. 154, 170–174, 1996. 147. Saarelainen, S., Seppala, E., Laasonen, K. and Hasan, J., Circulating endothelin-1 in obstructive sleep apnea, Endothelium 5, 115–118, 1997. 148. Schulz, R., Mahmoudi, S., Hattar, K., Sibelius, U., Olschewski, H., Mayer, K., Seeger, W. and Grimminger, F., Enhanced release of superoxide from polymorphonuclear neutrophils in obstructive sleep apnea. Impact of continuous positive airway pressure therapy, Am. J. Respir. Crit. Care Med. 162, 566–570, 2000. 149. Schulz, R., Schmidt, D., Blum, A., Lopes-Ribeiro, X., Lucke, C., Mayer, K., Olschewski, H., Seeger, W. and Grimminger, F., Decreased plasma levels of nitric oxide derivatives in obstructive sleep apnoea: Response to CPAP therapy, Thorax 55, 1046–1051, 2000. 150. Dyugovskaya, L., Lavie, P. and Lavie, L., Increased adhesion molecules expression and production of reactive oxygen species in leukocytes of sleep apnea patients, Am. J. Respir. Crit. Care Med. 165, 934–939, 2002. 151. O’Donnell, C.P., Allan, L., Atkinson, P. and Schwartz, A.R., The effect of upper airway obstruction and arousal on peripheral arterial tonometry in obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 166, 965–971, 2002. 152. Sullivan, C.E., Issa, F.G., Berthon-Jones, M. and Eves, L., Reversal of obstructive sleep apnoea by continuous positive airway pressure applied through the nares, Lancet 1, 862–865, 1981. 153. Sullivan, C.E., Berthon-Jones, M. and Issa, F.G., Remission of severe obesityhypoventilation syndrome after short-term treatment during sleep with nasal continuous positive airway pressure, Am. Rev. Respir. Dis. 128, 177–181. 154. McEvoy, R.D. and Thornton, A.T., Treatment of obstructive sleep apnea syndrome with nasal continuous positive airway pressure, Sleep 7, 313–325, 1984. 155. Frith, R.W. and Cant, B.R., Severe obstructive sleep apnoea treated with long term nasal continuous positive airway pressure, Thorax 40, 45–50, 1985.
Upper Airway Obstruction
347
156. Sanders, M.H. and Kern, N., Obstructive sleep apnea treated by independently adjusted inspiratory and expiratory positive airway pressures via nasal mask. Physiologic and clinical implications, Chest 98, 317–324, 1990. 157. Coccagna, G., Mantovani, M., Brignani, F., Parchi, C. and Lugaresi, E., Tracheostomy in hypersomnia with periodic breathing, Bull. Physiopathol. Respir. (Nancy) 8, 1217–1227, 1972. 158. Weitzman, E.D., Kahn, E. and Pollak, C.P., Quantitative analysis of sleep and sleep apnea before and after tracheostomy in patients with the hypersomnia–sleep apnea syndrome, Sleep 3, 407–423, 1980. 159. Safar, P., Escarraga, L.S. and Chang, F., Upper airway obstruction in the unconscious patient, J. Appl. Physiol. 14, 760–764, 1959. 160. Morikawa, S., Safar, P. and DeCarlo, J., Influence of the head-jaw position upon upper airway patency, Anesthesiology 22, 265–270, 1961. 161. Suratt, P.M., Dee, P., Atkinson, R.L., Armstrong, P. and Wilhoit, S.C., Fluoroscopic and computed tomographic features of the pharyngeal airway in obstructive sleep apnea, Am. Rev. Respir. Dis. 127, 487–492, 1983. 162. Smith, T.H., Baska, R.E., Francisco, C.B., McCray, G.M. and Kunz, S., Sleep apnea syndrome: Diagnosis of upper airway obstruction by fluoroscopy, J. Pediatr. 93, 891–892, 1978. 163. Shepard, J.W.J. and Thawley, S.E., Localization of upper airway collapse during sleep in patients with obstructive sleep apnea, Am. Rev. Respir. Dis. 141, 1350–1355, 1990. 164. Hudgel, D.W., Variable site of airway narrowing among obstructive sleep apnea patients, J. Appl. Physiol. 61, 1403–1409, 1986. 165. Boudewyns, A., Van de Heyning, P.H. and De Backer, W.A., Site of upper airway obstruction in obstructive apnoea and influence of sleep stage, Eur. Respir. J. 10, 2566–2572, 1997. 166. Shepard, J.W.J. and Thawley, S.E., Evaluation of the upper airway by computerized tomography in patients undergoing uvulopalatopharyngoplasty for obstructive sleep apnea, Am. Rev. Respir. Dis. 140, 711–716, 1989. 167. Isono, S., Remmers, J.E., Tanaka, A., Sho, Y., Sato, J. and Nishino, T., Anatomy of pharynx in patients with obstructive sleep apnea and in normal subjects, J. Appl. Physiol. 82, 1319–1326, 1997. 168. Badr, M.S., Toiber, F., Skatrud, J.B. and Dempsey, J., Pharyngeal narrowing/occlusion during central sleep apnea, J. Appl. Physiol. 78, 1806–1815, 1995. 169. Schneider, H., Boudewyns, A., Smith, P.L., O’Donnell, C.P., Canisius, S., Stammnitz, A., Allan, L. and Schwartz, A.R., Modulation of upper airway collapsibility during sleep: Influence of respiratory phase and flow regimen, J. Appl. Physiol. 93, 1365–1376, 2002. 170. Permutt, S. and Riley, R.L., Hemodynamics of collapsible vessels with tone: The vascular waterfall, J. Appl. Physiol. 18, 924–932, 1963. 171. Pride, N.B., Permutt, S., Riley, R.L. and Bromberger-Barnea, B., Determinants of maximal expiratory flow from the lungs, J. Appl. Physiol. 23, 646–662, 1967. 172. Lambert, R.K. and Wilson, T.A., Flow limitation in a collapsible tube, J. Appl. Physiol. 33, 150–153, 1972.
348
Patil et al.
173. Schwartz, A.R., Smith, P.L., Wise, R.A., Bankman, I. and Permutt, S., Effect of positive nasal pressure on upper airway pressure-flow relationships, J. Appl. Physiol. 66, 1626–1634, 1989. 174. Gold, A.R. and Schwartz, A.R., The pharyngeal critical pressure. The whys and hows of using nasal continuous positive airway pressure diagnostically, Chest 110, 1077–1088, 1996. 175. Schwartz, A.R., Schubert, N., Rothman, W., Godley, F., Marsh, B., Eisele, D., Nadeau, J., Permutt, L., Gleadhill, I. and Smith, P.L., Effect of uvulopalatopharyngoplasty on upper airway collapsibility in obstructive sleep apnea, Am. Rev. Respir. Dis. 145, 527–532, 1992. 176. Boudewyns, A., Punjabi, N., Van de Heyning, P.H., De Backer, W.A., O’Donnell, C.P., Schneider, H., Smith, P.L. and Schwartz, A.R., Abbreviated method for assessing upper airway function in obstructive sleep apnea, Chest 118, 1031–1041, 2000. 177. Penzel, T., Moller, M., Becker, H.F., Knaack, L. and Peter, J.H., Effect of sleep position and sleep stage on the collapsibility of the upper airways in patients with sleep apnea, Sleep 24, 90–95, 2001. 178. Neill, A.M., Angus, S.M., Sajkov, D. and McEvoy, R.D., Effects of sleep posture on upper airway stability in patients with obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 155, 199–204, 1997. 179. Farre, R., Peslin, R., Montserrat, J.M., Rotger, M. and Navajas, D., Flow-dependent positive airway pressure to maintain airway patency in sleep apnea–hypopnea syndrome, Am. J. Respir. Crit. Care Med. 157, 1855–1863, 1998. 180. King, E.D., O’Donnell, C.P., Smith, P.L. and Schwartz, A.R., A model of obstructive sleep apnea in normal humans. Role of the upper airway, Am. J. Respir. Crit. Care Med. 161, 1979–1984, 2000. 181. Isono, S., Morrison, D.L., Launois, S.H., Feroah, T.R., Whitelaw, W.A. and Remmers, J.E., Static mechanics of the velopharynx of patients with obstructive sleep apnea, J. Appl. Physiol. 75, 148–154, 1993. 182. Morrison, D.L., Launois, S.H., Isono, S., Feroah, T.R., Whitelaw, W.A. and Remmers, J.E., Pharyngeal narrowing and closing pressures in patients with obstructive sleep apnea, Am. Rev. Respir. Dis. 148, 606–611, 1993. 183. Isono, S., Feroah, T.R., Hajduk, E.A., Brant, R., Whitelaw, W.A. and Remmers, J.E., Interaction of cross-sectional area, driving pressure, and airflow of passive velopharynx, J. Appl. Physiol. 83, 851–859, 1997. 184. Isono, S., Shimada, A., Utsugi, M., Konno, A. and Nishino, T., Comparison of static mechanical properties of the passive pharynx between normal children and children with sleep-disordered breathing, Am. J. Respir. Crit. Care Med. 157, 1204–1212, 1998. 185. Isono, S., Tanaka, A., Sho, Y., Konno, A. and Nishino, T., Advancement of the mandible improves velopharyngeal airway patency, J. Appl. Physiol. 79, 2132–2138, 1995. 186. Launois, S.H., Feroah, T.R., Campbell, W.N., Issa, F.G., Morrison, D., Whitelaw, W.A., Isono, S. and Remmers, J.E., Site of pharyngeal narrowing predicts outcome of surgery for obstructive sleep apnea, Am. Rev. Respir. Dis. 147, 182–189, 1993.
Upper Airway Obstruction
349
187. Isono, S., Tanaka, A., Tagaito, Y., Sho, Y. and Nishino, T., Pharyngeal patency in response to advancement of the mandible in obese anesthetized persons, Anesthesiology 87, 1055–1062, 1997. 188. Isono, S., Saeki, N., Tanaka, A. and Nishino, T., Collapsibility of passive pharynx in patients with acromegaly, Am. J. Respir. Crit. Care Med. 160, 64–68, 1999. 189. Kato, J., Isono, S., Tanaka, A., Watanabe, T., Araki, D., Tanzawa, H. and Nishino, T., Dose-dependent effects of mandibular advancement on pharyngeal mechanics and nocturnal oxygenation in patients with sleep-disordered breathing, Chest 117, 1065–1072, 2000. 190. Thut, D.C., Schwartz, A.R., Roach, D., Wise, R.A., Permutt, S. and Smith, P.L., Tracheal and neck position influence upper airway airflow dynamics by altering airway length, J. Appl. Physiol. 75, 2084–2090, 1993. 191. Rowley, J.A., Permutt, S., Willey, S., Smith, P.L. and Schwartz, A.R., Effect of tracheal and tongue displacement on upper airway airflow dynamics, J. Appl. Physiol. 80, 2171–2178, 1996. 192. Van de Graaff, W.B., Thoracic influence on upper airway patency, J. Appl. Physiol. 65, 2124–2131, 1988. 193. Van de Graaff, W.B., Thoracic traction on the trachea: Mechanisms and magnitude, J. Appl. Physiol. 70, 1328–1336, 1991. 194. Series, F. and Marc, I., Effects of continuous negative airway pressurerelated lung deflation on upper airway collapsibility, J. Appl. Physiol. 75, 1222–1225, 1993. 195. Koenig, J.S. and Thach, B.T., Effects of mass loading on the upper airway, J. Appl. Physiol. 64, 2294–2299, 1988. 196. Katz, I., Stradling, J., Slutsky, A.S., Zamel, N. and Hoffstein, V., Do patients with obstructive sleep apnea have thick necks? Am. Rev. Respir. Dis. 141, 1228–1231, 1990. 197. Davies, R.J. and Stradling, J.R., The relationship between neck circumference, radiographic pharyngeal anatomy, and the obstructive sleep apnoea syndrome, Eur. Respir. J. 3, 509–514, 1990. 198. Shelton, K.E., Woodson, H., Gay, S. and Suratt, P.M., Pharyngeal fat in obstructive sleep apnea, Am. Rev. Respir. Dis. 148, 462–466, 1993. 199. Stauffer, J.L., Buick, M.K., Bixler, E.O., Sharkey, F.E., Abt, A.B., Manders, E.K., Kales, A., Cadieux, R.J., Barry, J.D. and Zwillich, C.W., Morphology of the uvula in obstructive sleep apnea, Am. Rev. Respir. Dis. 140, 724–728, 1989. 200. Sharp, J.T., Henry, J.P., Sweany, S.K., Meadows, W.R. and Pietras, R.J., Effects of mass loading the respiratory system in man, J. Appl. Physiol. 19, 959–966, 1964. 201. Series, F. and Marc, I., Influence of lung volume dependence of upper airway resistance during continuous negative airway pressure, J. Appl. Physiol. 77, 840–844, 1994. 202. Series, F., Cormier, Y. and Desmeules, M., Influence of passive changes of lung volume on upper airways, J. Appl. Physiol. 68, 2159–2164, 1990. 203. Eastwood, P.R., Szollosi, I., Platt, P.R. and Hillman, D.R., Comparison of upper airway collapse during general anaesthesia and sleep, Lancet 359, 1207–1209, 2002.
350
Patil et al.
204. Eastwood, P.R., Szollosi, I., Platt, P.R. and Hillman, D.R., Collapsibility of the upper airway during anesthesia with isoflurane, Anesthesiology 97, 786–793, 2002. 205. Mezzanotte, W.S., Tangel, D.J. and White, D.P., Waking genioglossal electromyogram in sleep apnea patients versus normal controls (a neuromuscular compensatory mechanism), J. Clin. Invest. 89, 1571–1579, 1992. 206. Petrof, B.J., Pack, A.I., Kelly, A.M., Eby, J. and Hendricks, J.C., Pharyngeal myopathy of loaded upper airway in dogs with sleep apnea, J. Appl. Physiol. 76, 1746–1752, 1994. 207. Hendricks, J.C., Petrof, B.J., Panckeri, K. and Pack, A.I., Upper airway dilating muscle hyperactivity during non-rapid eye movement sleep in English bulldogs, Am. Rev. Respir. Dis. 148, 185–194, 1993. 208. Series, F., Cote, C., Simoneau, J.A., Gelinas, Y., St Pierre, S., Leclerc, J., Ferland, R. and Marc, I., Physiologic, metabolic, and muscle fiber type characteristics of musculus uvulae in sleep apnea hypopnea syndrome and in snorers, J. Clin. Invest. 95, 20–25, 1995. 209. Remmers, J.E. and Bartlett, D.J., Reflex control of expiratory airflow and duration, J. Appl. Physiol. 42, 80–87, 1977. 210. Van Lunteren, E., Van de Graaff, W.B., Parker, D.M., Mitra, J., Haxhiu, M.A., Strohl, K.P. and Cherniack, N.S., Nasal and laryngeal reflex responses to negative upper airway pressure, J. Appl. Physiol. 56, 746–752, 1984. 211. Brouillette, R.T. and Thach, B.T., A neuromuscular mechanism maintaining extrathoracic airway patency, J. Appl. Physiol. 46, 772–779, 1979. 212. Weiner, D., Mitra, J., Salamone, J. and Cherniack, N.S., Effect of chemical stimuli on nerves supplying upper airway muscles, J. Appl. Physiol. 52, 530–536, 1982. 213. Rowley, J.A., Williams, B.C., Smith, P.L. and Schwartz, A.R., Neuromuscular activity and upper airway collapsibility. Mechanisms of action in the decerebrate cat, Am. J. Respir. Crit. Care Med. 156, 515–521, 1997. 214. Sant’Ambrogio, G., Tsubone, H. and Sant’Ambrogio, F.B., Sensory information from the upper airway: Role in the control of breathing, Respir. Physiol. 102, 1–16, 1995. 215. Mathew, O.P., Abu-Osba, Y.K. and Thach, B.T., Genioglossus muscle responses to upper airway pressure changes: Afferent pathways, J. Appl. Physiol. 52, 445–450, 1982. 216. Mathew, O.P., Abu-Osba, Y.K. and Thach, B.T., Influence of upper airway pressure changes on genioglossus muscle respiratory activity, J. Appl. Physiol. 52, 438–444, 1982. 217. Kuna, S.T., Interaction of hypercapnia and phasic volume feedback on motor control of the upper airway, J. Appl. Physiol. 63, 1744–1749, 1987. 218. Seelagy, M.M., Schwartz, A.R., Russ, D.B., King, E.D., Wise, R.A. and Smith, P.L., Reflex modulation of airflow dynamics through the upper airway, J. Appl. Physiol. 76, 2692–2700, 1994. 219. Schwartz, A.R., Thut, D.C., Brower, R.G., Gauda, E.B., Roach, D., Permutt, S. and Smith, P.L., Modulation of maximal inspiratory airflow by neuromuscular activity: Effect of CO2, J. Appl. Physiol. 74, 1597–1605, 1993.
Upper Airway Obstruction
351
220. Shea, S.A., Akahoshi, T., Edwards, J.K. and White, D.P., Influence of chemoreceptor stimuli on genioglossal response to negative pressure in humans, Am. J. Respir. Crit. Care Med. 162, 559–565, 2000. 221. Pillar, G., Malhotra, A., Fogel, R.B., Beauregard, J., Slamowitz, D.I., Shea, S.A. and White, D.P., Upper airway muscle responsiveness to rising PCO2 during NREM sleep, J. Appl. Physiol. 89, 1275–1282, 2000. 222. Mathew, O.P., Upper airway negative-pressure effects on respiratory activity of upper airway muscles, J. Appl. Physiol. 56, 500–505, 1984. 223. Aronson, R.M., Onal, E., Carley, D.W. and Lopata, M., Upper airway and respiratory muscle responses to continuous negative airway pressure, J. Appl. Physiol. 66, 1373–1382, 1989. 224. Sanna, A., Veriter, C. and Stanescu, D., Upper airway obstruction induced by negative-pressure ventilation in awake healthy subjects, J. Appl. Physiol. 75, 546–552, 1993. 225. Kuna, S.T., Insalaco, G. and Woodson, G.E., Thyroarytenoid muscle activity during wakefulness and sleep in normal adults, J. Appl. Physiol. 65, 1332–1339, 1988. 226. Wheatley, J.R., Tangel, D.J., Mezzanotte, W.S. and White, D.P., Influence of sleep on response to negative airway pressure of tensor palatini muscle and retropalatal airway, J. Appl. Physiol. 75, 2117–2124, 1993. 227. Sanna, A., Veriter, C., Kurtansky, A. and Stanescu, D., Contraction and relaxation of upper airway muscles during expiratory application of negative pressure at the mouth, Sleep 17, 220–225, 1994. 228. McNicholas, W.T., Coffey, M., McDonnell, T., O’Regan, R. and Fitzgerald, M.X., Upper airway obstruction during sleep in normal subjects after selective topical oropharyngeal anesthesia, Am. Rev. Respir. Dis. 135, 1316–1319, 1987. 229. Chadwick, G.A., Crowley, P., Fitzgerald, M.X., O’Regan, R.G. and McNicholas, W.T., Obstructive sleep apnea following topical oropharyngeal anesthesia in loud snorers, Am. Rev. Respir. Dis. 143, 810–813, 1991. 230. Berry, R.B., Kouchi, K.G., Bower, J.L. and Light, R.W., Effect of upper airway anesthesia on obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 151, 1857–1861, 1995. 231. Deegan, P.C., Mulloy, E. and McNicholas, W.T., Topical oropharyngeal anesthesia in patients with obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 151, 1108–1112, 1995. 232. Larsson, H., Carlsson-Nordlander, B., Lindblad, L.E., Norbeck, O. and Svanborg, E., Temperature thresholds in the oropharynx of patients with obstructive sleep apnea syndrome, Am. Rev. Respir. Dis. 146, 1246–1249, 1992. 233. Kimoff, R.J., Sforza, E., Champagne, V., Ofiara, L. and Gendron, D., Upper airway sensation in snoring and obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 164, 250–255, 2001. 234. Guilleminault, C., Li, K., Chen, N.H. and Poyares, D., Two-point palatal discrimination in patients with upper airway resistance syndrome, obstructive sleep apnea syndrome, and normal control subjects, Chest 122, 866–870, 2002.
352
Patil et al.
235. Woodson, B.T., Garancis, J.C. and Toohill, R.J., Histopathologic changes in snoring and obstructive sleep apnea syndrome, Laryngoscope 101, 1318–1322, 1991. 236. Edstrom, L., Larsson, H. and Larsson, L., Neurogenic effects on the palatopharyngeal muscle in patients with obstructive sleep apnoea: A muscle biopsy study, J. Neurol. Neurosurg. Psychiatry 55, 916–920, 1992. 237. Friberg, D., Gazelius, B., Hokfelt, T. and Nordlander, B., Abnormal afferent nerve endings in the soft palatal mucosa of sleep apnoics and habitual snorers, Regul. Pept. 71, 29–36, 1997. 238. Friberg, D., Ansved, T., Borg, K., Carlsson-Nordlander, B., Larsson, H. and Svanborg, E., Histological indications of a progressive snorers disease in an upper airway muscle, Am. J. Respir. Crit. Care Med. 157, 586–593, 1998. 239. Lindman, R. and Stal, P.S., Abnormal palatopharyngeal muscle morphology in sleep-disordered breathing, J. Neurol. Sci. 195, 11–23, 2002. 240. Malhotra, A., Fogel, R.B., Edwards, J.K., Shea, S.A. and White, D.P., Local mechanisms drive genioglossus activation in obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 161, 1746–1749, 2000. 241. Akahoshi, T., White, D.P., Edwards, J.K., Beauregard, J. and Shea, S.A., Phasic mechanoreceptor stimuli can induce phasic activation of upper airway muscles in humans, J. Physiol. 531, 677–691, 2001. 242. Miki, H., Hida, W., Shindoh, C., Kikuchi, Y., Chonan, T., Taguchi, O., Inoue, H. and Takishima, T., Effects of electrical stimulation of the genioglossus on upper airway resistance in anesthetized dogs, Am. Rev. Respir. Dis. 140, 1279–1284, 1989. 243. Bishara, H., Odeh, M., Schnall, R.P., Gavriely, N. and Oliven, A., Electrically-activated dilator muscles reduce pharyngeal resistance in anaesthetized dogs with upper airway obstruction, Eur. Respir. J. 8, 1537–1542, 1995. 244. Oliven, A., Odeh, M. and Schnall, R.P., Improved upper airway patency elicited by electrical stimulation of the hypoglossus nerves, Respiration 63, 213–216, 1996. 245. Schwartz, A.R., Thut, D.C., Russ, B., Seelagy, M., Yuan, X., Brower, R.G., Permutt, S., Wise, R.A. and Smith, P.L., Effect of electrical stimulation of the hypoglossal nerve on airflow mechanics in the isolated upper airway, Am. Rev. Respir. Dis. 147, 1144–1150, 1993. 246. Eisele, D.W., Schwartz, A.R., Hari, A., Thut, D.C. and Smith, P.L., The effects of selective nerve stimulation on upper airway airflow mechanics, Arch. Otolaryngol. Head Neck Surg. 121, 1361–1364, 1995. 247. McWhorter, A.J., Rowley, J.A., Eisele, D.W., Smith, P.L. and Schwartz, A.R., The effect of tensor veli palatini stimulation on upper airway patency, Arch. Otolaryngol. Head Neck Surg. 125, 937–940, 1999. 248. Fuller, D., Williams, J.S., Janssen, P.L. and Fregosi, R.F., Effect of co-activation of tongue protrudor and retractor muscles on tongue movements and pharyngeal airflow mechanics in the rat, J. Physiol. (Lond.) 519: Pt 2, 601–613, 1999.
Upper Airway Obstruction
353
249. Fuller, D., Mateika, J.H. and Fregosi, R.F., Co-activation of tongue protrudor and retractor muscles during chemoreceptor stimulation in the rat, J. Physiol. 507, 265–276, 1998. 250. Fuller, D.D. and Fregosi, R.F., Fatiguing contractions of tongue protrudor and retractor muscles: Influence of systemic hypoxia, J. Appl. Physiol. 88, 2123–2130, 2000. 251. Miki, H., Hida, W., Chonan, T., Kikuchi, Y. and Takishima, T., Effects of submental electrical stimulation during sleep on upper airway patency in patients with obstructive sleep apnea, Am. Rev. Respir. Dis. 140, 1285–1289, 1989. 252. Schwartz, A.R., Eisele, D.W., Hari, A., Testerman, R., Erickson, D. and Smith, P.L., Electrical stimulation of the lingual musculature in obstructive sleep apnea, J. Appl. Physiol. 81, 643–652, 1996. 253. Eisele, D.W., Smith, P.L., Alam, D.S. and Schwartz, A.R., Direct hypoglossal nerve stimulation in obstructive sleep apnea, Arch. Otolaryngol. Head Neck Surg. 123, 57–61, 1997. 254. Schwartz, A.R., Bennett, M.L., Smith, P.L., De Backer, W., Hedner, J., Boudewyns, A., Van de Heyning, P., Ejnell, H., Hochban, W., Knaack, L., Podszus, T., Penzel, T., Peter, J.H., Goding, G.S., Erickson, D.J., Testerman, R., Ottenhoff, F. and Eisele, D.W., Therapeutic electrical stimulation of the hypoglossal nerve in obstructive sleep apnea, Arch. Otolaryngol. Head Neck Surg. 127, 1216–1223, 2001. 255. Odeh, M., Schnall, R., Gavriely, N. and Oliven, A., Effect of upper airway muscle contraction on supraglottic resistance and stability, Respir. Physiol. 92, 139–150, 1993. 256. Odeh, M., Schnall, R., Gavriely, N. and Oliven, A., Dependency of upper airway patency on head position: The effect of muscle contraction, Respir. Physiol. 100, 239–244, 1995. 257. Bishara, H., Odeh, M., Schnall, R.P., Gavriely, N. and Oliven, A., Electrically-activated dilator muscles reduce pharyngeal resistance in anaesthetized dogs with upper airway obstruction, Eur. Respir. J. 8, 1537–1542, 1995. 258. Oliven, A., Schnall, R.P., Pillar, G., Gavriely, N. and Odeh, M., Sublingual electrical stimulation of the tongue during wakefulness and sleep, Respir. Physiol. 127, 217–226, 2001. 259. Schnall, R.P., Pillar, G., Kelsen, S.G. and Oliven, A., Dilatory effects of upper airway muscle contraction induced by electrical stimulation in awake humans, J. Appl. Physiol. 78, 1950–1956, 1995. 260. Kuna, S.T. and Vanoye, C.R., Respiratory-related pharyngeal constrictor muscle activity in decerebrate cats, J. Appl. Physiol. 83, 1588–1594, 1997. 261. Kuna, S.T. and Vanoye, C.R., Mechanical effects of pharyngeal constrictor activation on pharyngeal airway function, J. Appl. Physiol. 86, 411–417, 1999. 262. O’Halloran, K.D., Herman, J.K. and Bisgard, G.E., Respiratory-related pharyngeal constrictor muscle activity in awake goats, Respir. Physiol. 116, 9–23, 1999.
354
Patil et al.
263. Adachi, T., Umezaki, T., Matsuse, T. and Shin, T., Changes in laryngeal muscle activities during hypercapnia in the cat. Otolaryngol. Head Neck Surg. 118, 537–544, 1998. 264. Kuna, S.T., Effects of pharyngeal muscle activation on airway size and configuration, Am. J. Respir. Crit. Care Med. 164, 1236–1241, 2001. 265. Kuna, S.T. and Brennick, M.J., Effects of pharyngeal muscle activation on airway pressure–area relationships, Am. J. Respir. Crit. Care Med. 166, 972–977, 2002. 266. Van Lunteren, E., Haxhiu, M.A. and Cherniack, N.S., Relation between upper airway volume and hyoid muscle length, J. Appl. Physiol. 63, 1443–1449, 1987. 267. Smith, P.L., O’Donnell, C.P., Allan, L. and Schwartz, A.R., A physiologic comparison of nasal and oral positive airway pressure, Chest 123, 689–694, 2003. 268. Rubinstein, I., Colapinto, N., Rotstein, L.E., Brown, I.G. and Hoffstein, V., Improvement in upper airway function after weight loss in patients with obstructive sleep apnea, Am. Rev. Respir. Dis. 138, 1192–1195, 1988. 269. Suratt, P.M., McTier, R.F., Findley, L.J., Pohl, S.L. and Wilhoit, S.C., Effect of very-low-calorie diets with weight loss on obstructive sleep apnea, Am. J. Clin. Nutr. 56, 182S–184S, 1992. 270. Fujita, S., UPPP for sleep apnea and snoring, Ear Nose Throat J. 63, 227–235, 1984. 271. Woodson, B.T. and Toohill, R.J., Transpalatal advancement pharyngoplasty for obstructive sleep apnea, Laryngoscope 103, 269–276, 1993. 272. Houghton, D.J., Camilleri, A.E. and Stone, P., Adult obstructive sleep apnoea syndrome and tonsillectomy, J. Laryngol. Otol. 111, 829–832, 1997. 273. Riley, R.W., Powell, N.B., Guilleminault, C. and Nino-Murcia, G., Maxillary, mandibular, and hyoid advancement: An alternative to tracheostomy in obstructive sleep apnea syndrome, Otolaryngol. Head Neck Surg. 94, 584–588, 1986. 274. Riley, R.W., Powell, N.B. and Guilleminault, C., Maxillary, mandibular, and hyoid advancement for treatment of obstructive sleep apnea: A review of 40 patients, J. Oral Maxillofac. Surg. 48, 20–26, 1990. 275. Riley, R.W., Powell, N.B. and Guilleminault, C., Obstructive sleep apnea syndrome: A review of 306 consecutively treated surgical patients, Otolaryngol. Head Neck Surg. 108, 117–125, 1993. 276. Riley, R.W., Powell, N.B. and Guilleminault, C., Obstructive sleep apnea and the hyoid: A revised surgical procedure, Otolaryngol. Head Neck Surg. 111, 717–721, 1994. 277. Hida, W., Okabe, S., Miki, H., Kikuchi, Y., Taguchi, O., Takishima, T. and Shirato, K., Effects of submental stimulation for several consecutive nights in patients with obstructive sleep apnoea, Thorax 49, 446–452, 1994. 278. Smith, P.L., Haponik, E.F., Allen, R.P. and Bleecker, E.R., The effects of protriptyline in sleep-disordered breathing, Am. Rev. Respir. Dis. 127, 8–13, 1983.
Upper Airway Obstruction
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279. Brownell, L.G., West, P., Sweatman, P., Acres, J.C. and Kryger, M.H., Protriptyline in obstructive sleep apnea: A double-blind trial, N. Engl. J. Med. 307, 1037–1042, 1982. 280. Conway, W.A., Zorick, F., Piccione, P. and Roth, T., Protriptyline in the treatment of sleep apnoea, Thorax 37, 49–53, 1982. 281. Clark, R.W., Schmidt, H.S., Schaal, S.F., Boudoulas, H. and Schuller, D.E., Sleep apnea: Treatment with protriptyline, Neurology 29, 1287–1292, 1979. 282. Gould, G.A., Whyte, K.F., Rhind, G.B., Airlie, M.A., Catterall, J.R., Shapiro, C.M. and Douglas, N.J., The sleep hypopnea syndrome, Am. Rev. Respir. Dis. 137, 895–898, 1988. 283. Hanzel, D.A., Proia, N.G. and Hudgel, D.W., Response of obstructive sleep apnea to fluoxetine and protriptyline, Chest 100, 416–421, 1991. 284. Kraiczi, H., Hedner, J., Dahlof, P., Ejnell, H. and Carlson, J., Effect of serotonin uptake inhibition on breathing during sleep and daytime symptoms in obstructive sleep apnea, Sleep 22, 61–67, 1999. 285. Berry, R.B., Yamaura, E.M., Gill, K. and Reist, C., Acute effects of paroxetine on genioglossus activity in obstructive sleep apnea, Sleep 22, 1087–1092, 1999. 286. Sunderram, J., Parisi, R.A. and Strobel, R.J., Serotonergic stimulation of the genioglossus and the response to nasal continuous positive airway pressure, Am. J. Respir. Crit. Care Med. 162, 925–929, 2000. 287. Veasey, S.C., Chachkes, J., Fenik, P. and Hendricks, J.C., The effects of ondansetron on sleep-disordered breathing in the English bulldog, Sleep 24, 155–160, 2001. 288. Veasey, S.C., Serotonin. Culprit or promising therapy for obstructive sleep apnea? Am. J. Respir. Crit. Care Med. 163, 1045–1047, 2001. 289. Veasey, S.C., Panckeri, K.A., Hoffman, E.A., Pack, A.I. and Hendricks, J.C., The effects of serotonin antagonists in an animal model of sleep-disordered breathing, Am. J. Respir. Crit. Care Med. 153, 776–786, 1996. 290. Veasey, S.C., Fenik, P., Panckeri, K., Pack, A.I. and Hendricks, J.C., The effects of trazodone with L-tryptophan on sleep-disordered breathing in the English bulldog, Am. J. Respir. Crit. Care Med. 160, 1659–1667, 1999. 291. Bonora, M., Shields, G.I., Knuth, S.L., Bartlett, D.J., Jr., St. John, W.M., Selective depression by ethanol of upper airway respiratory motor activity in cats, Am. Rev. Respir. Dis. 130, 156–161, 1984. 292. Krol, R.C., Knuth, S.L. and Bartlett, D.J., Selective reduction of genioglossal muscle activity by alcohol in normal human subjects, Am. Rev. Respir. Dis. 129, 247–250, 1984. 293. Leiter, J.C., Doble, E.A., Knuth, S.L. and Bartlett, D.J., Respiratory activity of genioglossus. Interaction between alcohol and the menstrual cycle, Am. Rev. Respir. Dis. 135, 383–386, 1987. 294. Berry, R.B., Bonnet, M.H. and Light, R.W., Effect of ethanol on the arousal response to airway occlusion during sleep in normal subjects, Am. Rev. Respir. Dis. 145, 445–452, 1992. 295. Issa, F.G. and Sullivan, C.E., Alcohol, snoring and sleep apnea, J. Neurol. Neurosurg. Psychiatry 45, 353–359, 1982.
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Patil et al.
296. Litman, R.S., Hayes, J.L., Basco, M.G., Schwartz, A.R., Bailey, P.L. and Ward, D.S., Use of dynamic negative airway pressure (DNAP) to assess sedative-induced upper airway obstruction, Anesthesiology 96, 342–345, 2002. 297. Rowley, J.A., Zhou, X., Vergine, I., Shkoukani, M.A. and Badr, M.S., Influence of gender on upper airway mechanics: Upper airway resistance and Pcrit, J. Appl. Physiol. 91, 2248–2254, 2001. 298. Eastwood, P.R., Szollosi, I., Platt, P.R. and Hillman, D.R., Collapsibility of the upper airway during anesthesia with isoflurane, Anesthesiology 97, 786–793, 2002. 299. Martin, T.J. and Sanders, M.H., Chronic alveolar hypoventilation: A review for clinicians, Sleep 18, 617–634, 1995. 300. Winakur, S.J., Smith, P.L. and Schwartz, A.R., Pathophysiology and risk factors for obstructive sleep apnea, Sem. Respir. Crit. Care Med. 19, 99–112, 1998.
11 High Altitude
FRANK L. POWELL
PHILIP E. BICKLER
University of California, San Diego, La Jolla, California
University of California, San Francisco, California
I.
Introduction
Control of breathing at high altitude has been studied intensely for over a century. Although there has been a variety of theories to explain the deleterious effects of high altitude, it is now clear that the primary problem is a decrease in the oxygen partial pressure. The decrease in inspired PO2 during ascent to high altitude is the most common form of environmental hypoxia in humans and terrestrial animals, and the reflex increase in ventilation from arterial hypoxemia is the body’s first line of defense against decreased O2 supply. Hence, understanding the control of breathing at altitude provides the basis for understanding the normal physiological response to hypoxia. We understand the basic elements of the response but many fundamental questions remain, and these are currently under investigation with modern experimental techniques. Although hypoxia is the primary physiological challenge at high altitude, it is important to emphasize that O2 is not the only factor that affects the control of breathing at altitude. Specifically, CO2 and pH are also extremely important because they are both determined by the level 357
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of ventilation, and because they stimulate ventilation. Another important factor in the control of breathing at high altitude is the duration of hypoxic exposure. Ventilatory acclimatization to high altitude provides a useful model to study the changes that can occur in control of breathing during chronic hypoxia, and can provide important insights into the control of breathing under pathological conditions such as heart and lung disease. Finally, there are pathological changes at high altitude and these may involve problems with control of breathing. This chapter reviews recent progress in our understanding of the control of breathing in hypoxia at high altitude, how this changes during prolonged exposure to altitude, and how it may be involved in pathological responses to altitude. II.
Ventilatory Response to High Altitude
Decreasing PiO2 at altitude results in decreased PaO2 , which stimulates arterial chemoreceptors. The ventilatory response to PaO2 is called the hypoxic ventilatory response, or HVR, and is the reflex response to stimulation of arterial chemoreceptors, including the carotid and aortic bodies. Normally, the HVR is not very large until PaO2 falls to a level at which O2-hemoglobin saturation starts decreasing significantly and, in fact, ventilation is a linear function of arterial O2 saturation down to 70–80% [1]. However, this linear relationship is a coincidence and not a mechanistic explanation because arterial chemoreceptors respond only to O2 partial pressure, and not O2 content or saturation [2]. This chapter does not review the mechanisms of chemoreception in the carotid body arterial chemoreceptors, which have been covered elsewhere ([2], and Nurse, Chapter 1, this volume). Arterial PCO2 is generally the most important regulated variable in the control of breathing under resting conditions in normoxia. Even under conditions of hypoxia, the effects of PaCO2 are important and will determine the magnitude of the HVR, as illustrated in Figure 11.1. The lower response curve shows a normal HVR with PaCO2 decreasing as a result of hypoxic ventilatory stimulation. Such a poikilocapnic HVR would occur during exposure to progressively higher altitudes. As PaO2 decreases below 60 mmHg, the arterial chemoreceptors are stimulated and cause a small increase in ventilation. This decreases PaCO2 and, thereby, ventilatory drive from central and arterial chemoreceptor stimulation. The net effect on ventilation is the result of stimulation by hypoxia and inhibition from decreased PaCO2 (i.e., 37 mmHg). As PaO2 decreases more, hypoxic ventilatory stimulation increases, and this causes further reductions in PaCO2 . Ventilation remains a compromise between hypoxic stimulation and hypocapnic inhibition on the non-isocapnic hypoxic ventilatory response.
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Figure 11.1 The hypoxic ventilatory response (HVR) under normal non-isocapnic (or poikilocapnic) conditions, with PaCO2 decreasing as ventilation is stimulated, and under isocapnic conditions at the normal or elevated PaCO2 levels. The synergistic interaction between CO2 and hypoxia as stimulants of the arterial chemoreceptors means that ventilation during hypoxic conditions at altitude is a compromise between hypoxic stimulation and hypocapnic inhibition.
To rigorously quantify the ventilatory response to PO2, one needs to maintain PaCO2 constant, for example by increasing inspired PCO2. This is shown on the isocapnic HVR in Figure 11.1; note that this is steeper than the poikilocapnic HVR. Increasing PaCO2 further above normal reveals a synergistic, or multiplicative, interaction between PaO2 and PaCO2. This multiplicative interaction can be explained by the effects of PaO2 and PaCO2 on the carotid body chemoreceptors, i.e., the afferent activity from the chemoreceptors reflects the multiplicative nature of these stimuli [2]. During longer exposures to hypoxia at high altitude, there are further changes in the ventilatory response to O2 and CO2. This results in ventilatory acclimatization to hypoxia (VAH), which includes both persistent hyperventilation when normoxia is restored, and an increase in the isocapnic HVR (Figure 11.2). The persistent hyperventilation in normoxia, and resulting decrease in PaCO2, is one of the most robust measures of VAH. This apparent increase in CO2 sensitivity can be viewed as a change in the arterial PCO2 set point with chronic hypoxia. The increase in O2 sensitivity has been more controversial, and is often confused by the blunted HVR observed in some high altitude natives, or in residents of high altitude for many years (see Hypoxic Desensitization
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PaO2 (Torr) Figure 11.2 Isocapnic HVR in rats under control, normoxic conditions (squares) and after acclimatization to hypoxia (simulating 6,000 m). Ventilatory acclimatization includes (1) a persistent hyperventilation when normoxia is restored, and (2) an increase in the isocapnic HVR. PaCO2 was maintained at the level measured during 30% O2 breathing, which removed any O2-sensitive stimulation of arterial chemoreceptors, and was lower after acclimatization (32 Torr vs. 37 Torr before), reflecting a change in the arterial PCO2 set-point after chronic hypoxia (Data from Ref. 51).
below). However, it is now clear that the isocapnic HVR is increased by chronic hypoxia in humans and animals. A recent review of the literature found that conflicting reports could be explained by (a) complications from different time domains of the HVR that may arise with different protocols; (b) indices used to quantify the HVR, and (c) the effects of CO2 on the HVR [3]. All studies of the isocapnic HVR in humans after acclimatization to hypoxia have found significant increases in the HVR. The exact choice of PaCO2 for the isocapnic measurements did not appear to be critical. Hence, VAH in normal conscious humans includes increased O2 sensitivity and a change in the arterial PO2 set point. Next, we review the mechanisms of the HVR that are important during different times of exposure to hypoxia at altitude, and then the changes in ventilatory responses to CO2 during acclimatization. III.
Time Domains of the HVR
The HVR is not a single mechanism, but is a complex interplay between several distinct mechanisms operating in different time domains [4]. Among
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the features distinguishing these different mechanisms are: (a) the specific stimuli that elicit them (e.g., pattern and intensity of hypoxic exposure); (b) their time course (seconds to years); (c) the nature of the reflex response (tidal volume vs. frequency); (d) whether the response is excitatory or inhibitory to ventilation, and (e) their neurochemical basis. Some mechanisms are sufficiently long lasting to affect future ventilatory responses to hypoxia, indicating a degree of memory, or functional plasticity in the ventilatory control system. All of these mechanisms can interact to affect the ventilatory response to hypoxia at altitude, and produce the timedependent changes of ventilatory acclimatization described above. Many of the mechanisms have been discussed in more detail in other reviews [4–9], which are cited in place of original references when possible. A. Responses to Acute Hypoxia
Changes in ventilation (in both VT and fR) during or following a brief hypoxic exposure (2–5 min) are shown schematically in Figure 11.3. Although the primary stimulus to these changes is a decrease in PaO2
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Figure 11.3 Time domains of the HVR. Top panel—Ventilation, tidal volume and respiratory frequency during acute hypoxia. The acute HVR (AR) is followed by short-term potentiation (STP) of VT and short-term depression (STD) of fR. STP and STD are manifested at the termination of acute hypoxia also. Bottom panel—Ventilation during chronic hypoxia shows hypoxic ventilatory decline (HVD) after the acute HVR, then an increase with ventilatory acclimatization to hypoxia (VAH). Chronic hypoxia for years to a lifetime leads to hypoxic desensitization (HD), although the exact duration necessary for this is not known.
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acting on the arterial (peripheral) chemoreceptors, similar responses with similar time courses can be elicited in anesthetized, paralyzed mammals by electrical stimulation of the carotid sinus nerve. Within this relatively short time domain, at least three distinct mechanisms have been identified: the Acute Response, Short-Term Potentiation and Short-Term Depression. Acute HVR
As discussed above, the acute HVR is the immediate augmentation of ventilatory activity at the onset of hypoxia (within one breath of PaO2 changing at the carotid bodies), and the decrease in ventilatory activity at the termination of hypoxia (Figure 11.3). The acute HVR represents the effects of changes in arterial (peripheral) chemoreceptor afferent input to glutamatergic synapses in the nucleus of the solitary tract, NTS (cf. [7,10]). This synaptic input is gated, such that the immediate response to changes in afferent input depends on the phase of the ongoing respiratory cycle [6]. This is the mechanism responsible for the rapid increase in ventilation during acute exposure to altitude. Short-Term Potentiation (STP)
STP represents a further, progressive increase in ventilatory activity after the acute HVR that proceeds with a time course of many seconds up to one minute [6]. STP is also manifested as a progressive post-stimulation decline in ventilatory activity after carotid sinus nerve stimulation, with a slightly longer time constant of 1–2 minutes (Figure 11.3). Although STP is generally thought to be a reflection of the same mechanism during the on and off responses, this hypothesis has not been rigorously tested. STP was formerly referred to as afterdischarge, based on Sherrington’s concept of a prolonged neural discharge triggered by a brief stimulation [6]. STP has been demonstrated following brief hypoxia in human subjects while awake [11], asleep [12] and during exercise [13]. STP, which increases ventilatory drive and changes the normal level for arterial PCO2, has not been studied after acclimatization to hypoxia (see below). STP does not appear specific to carotid body afferent inputs since other respiratory stimuli elicit a similar phenomenon [6]. Several neurotransmitters and neuromodulators (e.g., serotonin, catecholamines, opiates) have been ruled out as candidates for mediating STP in mammals [6]. Possible explanations for STP include: (a) presynaptic calcium accumulation along premotor pathways, thus mediating enhanced transmitter release when action potentials subsequently reach the terminal [14], or (b) release of modulatory neuropeptides, such as substance P, at key locations in the respiratory neural control system.
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STP has been suggested to play a smoothing role in the control of breathing, preventing reflex activation of the respiratory system from proceeding too rapidly and, thus, imparting system stability [6]. Short-Term Depression (STD)
STD is manifested as the recovery of the transient overshoot in respiratory frequency at the onset of carotid chemoreceptor stimulation, or a transient undershoot in frequency at the termination of stimulation, that lasts many seconds to a few minutes (Figure 11.3). To date, STD has been demonstrated only in the fR response of phrenic nerve activity in anesthetized rats during or following hypoxia or carotid sinus nerve stimulation [15]. Although STD is similar in some respects to hypoxic ventilatory decline (HVD, see below), important differences lead us to classify this time-dependent response as a unique mechanism or class of mechanisms at this time. There are differences in the ventilatory pattern (fR decreases in STD vs. VT in HVD), time course (many seconds to a few minutes with STD vs. many minutes with HVD), and in the neurochemical mechanisms thought to be involved in STD and HVD. It is possible that STD is unique to anesthetized rats, since it is not observed in anesthetized cats following carotid sinus nerve stimulation (cf. [6]) nor in other mammalian species investigated (e.g., goats, [16]). B. Responses to Sustained Hypoxia
Additional mechanisms become apparent when the hypoxic exposure is continuous for a prolonged period of several minutes to months. Among these mechanisms are: Hypoxic Ventilatory Decline, Ventilatory Acclimatization and Deacclimatization to Hypoxia during chronic exposures, and Hypoxic Desensitization with life-long hypoxia (Figure 11.3). Hypoxic Ventilatory Decline (HVD)
HVD is the roll off, or decrease in ventilation relative to the acute HVR, when moderate hypoxemia is sustained for 5 to 30 minutes in adult animals (Figure 11.3) [5]. HVD is distinct from the secondary decrease in ventilation from hypocapnia accompanying the acute response because it occurs during isocapnic hypoxia. As defined here, HVD differs from the biphasic HVR observed in neonates and small rodents, which represents an appropriate decrease in ventilation for the decrease in metabolic rate that occurs with hypoxia in small animals. The onset and resolution of HVD have a similar time course. Once HVD is established, the ventilatory response to subsequent hypoxic challenges is depressed for up to 60 min after normoxia is restored, but this can be shortened by breathing O2 enriched gas mixtures [17].
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HVD has been observed in awake humans, and in awake and anesthetized cats, but it could not be demonstrated in awake dogs [5] and may, or may not, be observed in awake rats [3]. It primarily affects VT, but not rhythm generation, and differs from short-term depression (STD, see above), which decreases fR. The mechanism of HVD is unknown but experiments on different species and preparations suggest specific effects of hypoxia on both (a) the sensitivity of ventilation to O2 and (b) central ventilatory drive independent of any changes in O2 sensitivity. Experiments from several studies in awake humans indicate that O2 sensitivity decreases during HVD, and it has been hypothesized that HVD is a specific effect of hypoxia on arterial chemoreceptors [18]. Some of the strongest support for this theory comes from experiments comparing the ventilatory response with brief stimulation of arterial chemoreceptors by hypoxia vs. hypercapnia during sustained hypoxia [19]. However, it is difficult to quantify the carotid body contribution to the ventilatory response to CO2 in awake humans. Other studies of awake humans have found ventilatory decline during sustained hypoxia without a significant decrease in O2 sensitivity [20,21]. This is consistent with experiments on anesthetized animals which show that the ventilatory response to arterial chemoreceptor or carotid sinus nerve stimulation, (i.e., the gain or slope of the HVR) does not decrease during several minutes of inspired hypoxia or low levels of carbon monoxide inhalation [17]. It is also consistent with neural recordings from carotid body afferents in animals, which, with the possible exception of the rabbit, show no change in chemoreceptor activity over the time course of HVD [5]. These data suggest that HVD is a decline in central ventilatory drive independent of changes in O2 sensitivity of arterial chemoreceptors or ventilation. The neurochemical basis of HVD is not clear. There is experimental evidence implicating ventilatory inhibition by adenosine, GABA, and opioids, but none of these neuromodulators can completely explain HVD [5]. A dopaminergic mechanism may be involved in HVD in some species since haloperidol, a D2 dopamine receptor antagonist, eliminates ventilatory roll-off in awake and anesthetized cats [5], but not in awake humans [22]. D2 receptors in the carotid body are not involved in the decrease in chemoreceptor O2 sensitivity during HVD in humans [5]. One mechanism that is apparently not involved is insufficient energy substrate [17]. Acid-base shifts in the brain at the site of the chemoreceptors have been proposed to contribute to HVD, although evidence for this is not clear. With the acute hyperventilation induced by hypoxia, the fall in tissue PCO2 at the central chemoreceptors should lower the ventilatory stimulus. However, at least in the case when CO2 at the peripheral chemoreceptors remains isocapnic, HVD is still observed [23]. Another test of this concept could be made with the administration of a carbonic anhydrase inhibitor
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such as acetazolamide (DiamoxÕ ), which would eliminate the brain tissue alkalinity resulting from hyperventilation [24], and possibly eliminate the HVD. A role for lactic acid accumulation in the mechanism for HVD has also been suggested. Hypocapnia and hypoxia are synergistic in increasing lactic acid production in the brain [25]. Somewhat surprisingly, lactic acid production in brain cells may depress ventilation, even though acidity generally stimulates it (reviewed by Smith et al. in Ref. [23]). Cells in the ventral medulla chemosensitive zones produce an excess of lactic acid during hypoxia [26,27]. Intracellular acidification may lead to depression of neuronal excitability by hyperpolarizing the membrane potential (by altering the transcellular Hþ gradient, or by activation of acid-sensitive potassium channels) or by reducing neurotransmitter release via reductions in intracellular [Ca2þ]i. This effect decreases respiratory motor output, explaining the decrease in ventilation when compared with that predicted. Testing this hypothesis will require improved identification of chemosensitive respiratory neurons and methods to correlate pHi, [Ca2þ]i and neuronal output. A potential mechanism for the increase in ventilation during altitude acclimatization (described next) is a decrease in HVD with chronic hypoxia. However, HVD has been shown to persist during prolonged exposure to altitude [1]. Ventilatory Acclimatization to Hypoxia (VAH)
VAH is defined as the time-dependent increase in ventilation that occurs with chronic hypoxic exposures of several hours to months (Figure 11.3) [5,8]. Ventilatory acclimatization to high altitude is the classic example of VAH, and its physiological significance, in terms of increasing O2 delivery, is well known [28]. The time course of VAH is species-dependent, and can be complete after only 4 to 6 hrs in goats (in terms of no further changes in ventilation or PaCO2 relative to seven days of hypoxia), or may require more than 10 days in humans [5]. The rat has a time course similar to that in humans, and has been shown to be a good model for human VAH when corrections are made for changes in metabolic rate occurring during the first days of hypoxia [5]. Experiments on VAH usually study the effects of chronic hypoxia and hypocapnia, both of which occur in healthy subjects exposed to hypoxia. The effects of changing CO2 sensitivity are discussed separately below. However, recent experiments show that the isocapnic HVR also increases in humans and animals [3]. Changes in HVR with VAH
Changes in ventilatory and carotid body O2 sensitivity are now generally accepted mechanisms contributing to the increased HVR with VAH. This is
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consistent with the lack of VAH in carotid body-denervated animals, and the increase in the isocapnic HVR during VAH in awake animals and humans [3,5,29]. Animal studies show increased O2 sensitivity of carotid body chemoreceptors, although there are significant species differences. In goats, carotid body chemoreceptor afferent activity increases during 6 hrs of hypoxia, similar to the time course required for complete VAH in this species [5]. In cats, carotid body chemoreceptors increase O2 sensitivity after 48 hrs of hypoxia [30], although the situation after 2 wks of hypoxia is unclear (see Hypoxic Desensitization below). The increase in carotid body chemoreceptor O2 sensitivity can completely explain VAH in goats after 6 hrs and cats after 48 hrs of hypoxia. However, it is important to understand that complete VAH after only hours does not preclude different mechanisms turning off and on during longer time domains. Another mechanism that can increase the HVR with VAH is altered CNS processing of arterial chemoreceptor afferent input. The first evidence for this came from human studies showing an increase in the ventilatory response to arterial chemoreceptor stimulation with doxapram after acclimatization to altitude [31]. If chronic hypoxia does not change the effect of this drug on chemoreceptors, then the change with acclimatization could be explained by an increase in the CNS gain of the HVR that translates chemoreceptor afferent input into ventilatory output. The ventilatory response to arterial chemoreceptor stimulation with doxapram or NaCN also increases significantly after exposure to two or more days of hypoxia in rats and ponies (reviewed by Powell, Huey, and Dwinell [32]). However, the ventilatory response to NaCN is not increased significantly after four hours of hypoxia in goats, so increases in the CNS gain of the HVR may require exposure to hypoxia lasting days or more. The most conclusive evidence for chronic hypoxia increasing the CNS gain of the HVR comes from experiments on anesthetized rats [33]. These experiments produced graded and reproducible changes in arterial chemoreceptor afferent input by electrically stimulating the carotid sinus nerve at different frequencies and measuring ventilatory motor output as integrated phrenic nerve activity. After seven days of hypoxia, the CNS gain of the HVR increased significantly, primarily by increasing the ventilatory frequency response to chemoreceptor stimulation. It was hypothesized that the increased CNS gain of the HVR takes more than two days of hypoxic exposure, which is consistent with it not being observed in the shorter studies (see above). Dopamine (DA) was suggested as a likely candidate for increasing the CNS gain of the HVR based on a significant correlation between increases in ventilation and tyrosine hydroxylase (the rate-limiting enzyme for DA synthesis) in respiratory centers of the brain after chronic hypoxia [34]. Further, DA acting on D2 receptors (D2-R) in the brain increases the CNS gain of the HVR [35]. Pharmacological and transgenic studies in mice
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support an increased excitatory effect of DA acting at D2-receptors (D2-R) in the CNS contributing to ventilatory acclimatization to hypoxia. Acclimatized mice with the D2-R gene ‘knocked out’ [36] or D2-R blocked with a systemic antagonist [37] do not show a normal time-dependent increase in hypoxic ventilation, and the independent effects of D2-R in the carotid body cannot explain these results [35]. However, D2-R in the CNS cannot explain the increased CNS gain of the HVR in acclimatized rats [38,39]. In rats, at least, it appears that changes in dopaminergic modulation of the HVR function to optimize CNS sensitivity to afferent input from arterial chemoreceptors, as well as to O2 sensitivity of the carotid body [35]. Glutamate and GABA are also involved in the HVR but they have not been investigated after chronic hypoxia [32]. Nitric oxide (NO) exerts positive feedback on glutamate release in the respiratory centers where the arterial chemoreceptors synapse, and could be involved in increasing the CNS gain of the HVR also [32]. Serotonin is important for long-term facilitation of ventilation following repeated bouts of hypoxia, but this mechanism cannot explain ventilatory acclimatization to sustained hypoxia [7]. In summary, the neurochemical basis for the increased CNS gain of the HVR with ventilatory acclimatization to hypoxia remains unknown. Changes in ventilatory drive and CO2 sensitivity with VAH
From the early 1960s, acid-base changes associated with the hyperventilation accompanying acclimatization to high altitude have been proposed to contribute to the pattern of ventilatory changes. Mitchell and Loeschke [40] had recently identified pH-sensitive areas on the ventral surface of the medulla that stimulated ventilation. The group of Severinghaus and Mitchell [41] proposed that the acute respiratory alkalosis from the acute HVR restrained the increase in alveolar ventilation because of the alkalinity in the cerebrospinal fluid (CSF) bathing these central chemoreceptors. During acclimatization, compensatory responses were proposed to lower the pH in the CSF (for example, by active transport of HCO 3 out of CSF) and to facilitate a further increase in ventilation. The shifts in CSF acid-base balance could persist for a time after return to low elevation (or during administration of supplemental oxygen at high altitude), explaining the Houston-Riley enigma [42] of continued hyperventilation after return to sea level (see Ventilatory Deacclimatization to Hypoxia below). However, subsequent studies failed to confirm that CSF pH returned to normal with time at altitude [43,44]; in fact, the CSF pH most likely remains increased for a long period. A problem in providing evidence that pH or bicarbonate compensation in the CSF contributes to VAH could be that very small changes in CSF pH have large effects on ventilation. For example, the pH changes in CSF required to produce a 1L/min change in ventilation (sufficient to decrease PaCO2 2–3 mmHg) is only ca. 0.001 pH units, below ready detection.
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Further, decreases in CSF bicarbonate change the relationship between pH and PCO2 making changes in the PCO2/pH relationship even steeper. In their study of CSF pH and respiratory sensitivity to hypercapnia during altitude acclimatization, Crawford and Severinghaus [44] found that only a 0.003 difference of CSF pH (based on the slope of the hypercapnic ventilatory response) could explain the persistent hyperventilation during normoxia after three days at altitude. Another complication for testing this hypothesis is the fact that the chemosensory cells responding to increases in CO2 may be few in number [20] and distributed throughout the brain stem. Nattie [45] reviews data showing that at least five distinct, non-medullary, chemoreceptor areas respond to microdialysis with CO2 or micro injections of acetazolamide by increasing respiratory motor neuron output in awake or anesthetized animals. Also, the pH measured by central chemoreceptors remains controversial [46]. Hence, it has not been possible to demonstrate conclusively that a change in stimulus level of central chemoreceptors contributes to an increase in ventilation with VAH. However, the increased ventilatory drive observed during VAH behaves as if the set point for central CO2 receptors is decreased. Ventilatory Deacclimatization from Hypoxia (VDH)
When normoxia is acutely restored during, or after, chronic hypoxia, ventilation and ventilatory O2 sensitivity do not immediately return to control levels [5]. This persistent hyperventilation in normoxia after chronic hypoxia is termed VDH, and it decays with a time course similar to the time-dependent increase in ventilation with VAH. Given the similar time courses for VAH and VDH, it was generally thought that they were the same mechanism being turned on and off, respectively. However, recent studies have shown that VDH and VAH can be dissociated, and are probably separate mechanisms. For example, VDH does not occur after VAH in goats exposed to isocapnic hypoxia for 4 to 6 hrs, but it does occur when hypoxic exposure is accompanied by hypocapnia [5]. Differences in VDH with CO2 levels support the idea that changes in CO2-sensitive mechanisms explain persistent hyperventilation in normoxia after acclimatization, as discussed above. Increased O2 sensitivity of arterial chemoreceptors, or the CNS gain of the HVR, would not be expected to increase ventilation in normoxia when arterial chemoreceptor afferent activity is minimal. However, a recent study found that hypocapnia is not necessary for VDH in humans [29]. The authors postulated hyperventilation-induced hyperpnea and potential central effects of hypoxia might explain persistent hyperventilation when normoxia is restored after chronic hypoxia.
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Hypoxic Desensitization (HD)
When humans experience chronic hypoxia for years or a lifetime, the HVR becomes blunted. Ventilation in hypoxia is decreased relative to normal subjects acclimatized to altitude for shorter periods of time (Figure 11.3) and ventilatory sensitivity to PaO2 is decreased [8]. This may conserve energy, through reduced work of breathing, when other non-ventilatory modes of acclimatization (e.g., metabolic, vascular, hematological) have had time to occur [8,28]. HD is an acquired characteristic that increases with the level of altitude and time at altitude, but there is disagreement about its reversibility [8]. An interesting exception to the correlation between high altitude exposure and HD is provided by adult Tibetans residing at 3658 m since birth. These high-altitude natives do not have a blunted HVR, in contrast to Chinese born near sea level but who lived at altitude for years. However, Tibetan natives from 4400 m do show HD [47]. This suggests that both genetic and environmental factors contribute to HD. The effects of genetics and potential evolutionary adaptations to hypoxia at altitude in humans have been reviewed in a special issue of the journal High Altitude Medicine and Biology [48–50]. One of the difficulties in studying the physiological mechanisms of HD is the lack of suitable animal models. Cats show HD after two weeks at simulated altitudes of 5500 m, but it reverses quickly with return to normoxia, in contrast to humans [8]. Reports of a blunted HVR in chronically hypoxic rats can be explained by the effects of hypocapnia or anesthesia on an HVR that is actually increased by chronic hypoxia [51]. The profound changes in carotid body structure that occur with chronic hypoxia might be expected to alter chemoreceptor O2 sensitivity, but many of these morphological changes occur before HD [5], so their significance in HD is not clear. Increased dopaminergic inhibition at the carotid body may be involved [5]. Changes in both carotid body chemoreceptor O2 sensitivity and CNS gain of the HVR are reported to contribute to HD in cats [52], but these results are difficult to reconcile with another study which shows increased carotid body chemoreceptor O2 sensitivity in cats after the same period of acclimatization [53]. C. Intermittent Hypoxia
The physiological responses to intermittent hypoxia (IH) are clearly not the same as those to continuous hypoxia, although the effects of the pattern and dose of hypoxic exposure are not yet understood. Much of the basic research in this area has been motivated by the problem of sleep apnea at sea level, although there have been several studies of IH for physical training to improve athletic performance at low altitudes, high-altitude mountaineering, commuting to work at high altitude, and even comparative physiological models in nature (reviewed by Garcia and Powell, Ref. [54]).
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The problem of sleep apnea at altitude is considered separately, because here we review the changes in ventilatory control with IH and high-altitude mountaineering or work. IH has been an integral part of high-altitude mountaineering for years. Climbers ascend from low-altitude base camps to higher altitudes where they establish camps to stage for their summit attempts. Typically they return to base camp for more supplies and sleep. Over the days and weeks that are necessary to climb major peaks, this pattern of up and down results in IH exposures that are typically increasing in the degree of hypoxia. Although this pattern probably originated out of logistical necessity, it has been obvious for many years that climbers should sleep at as low an altitude as possible, while still obtaining adequate acclimatization and meeting the climbing schedule for a successful summit bid. More recently, IH has been viewed as a way to potentially speed up or improve acclimatization. The tragic events on Mount Everest in 1996 focused attention on IH as a means to gain rapid acclimatization. Traditionally, acclimatization in the Himalayas is achieved by slowly ascending to altitude during a long trek. In contrast, modern guided trips taking paying clients to the highest summit on earth have a strong profit motive to minimize the total time for an expedition. Of course, minimizing exposure to hypoxia is a reasonable goal but acclimatization cannot occur without some hypoxia. Several groups have studied the applied physiology of IH as a means of pre-acclimatization to improve climbing performance at high altitude. One of the earliest studies hypothesized that IH could produce acclimatization more efficiently than continuous chronic hypoxia (CH) [55]. To mimic the usual climbing pattern of mountaineers during an expedition, they exposed 12 subjects to simulated altitude in a hypobaric chamber. After three consecutive days simulating 6000 m for 5 hr and 8000 m for the next one hour, they observed an increase in VE and PaO2 in hypoxia, indicating the initiation of ventilatory acclimatization. Subsequent studies exposed resting humans to IH simulating altitudes above 5000 m as a pre-acclimatization training method for mountaineering expeditions [20,45,56–58]. Others used less severe hypoxic stimuli (simulated altitudes of 2500 to 5000 m) but added exercise to the IH and also measured the effects of such IH on the hypoxic ventilatory response (HVR) [59–62]. In general these studies showed increases in the HVR and ventilation and arterial saturation (SaO2) during hypoxia. Hence, the studies show ventilatory acclimatization to IH is qualitatively similar to CH, although they did not specifically compare the two patterns of hypoxia. Also, some of the protocols actually combined stimulus patterns by administering CH before IH [56,57]. The opportunities for work at high altitude are increasing with the extensive development of commercial and scientific activities in high
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mountain ranges. Notable examples include mining in South America and high-altitude observatories for optical and radio telescopes, both involving altitudes of 3500 to 6000 m [63–66]. With these opportunities come large-scale challenges to public health and worker safety issues, as well as problems in maintaining worker productivity and morale. Not only are there physical effects on the employees from working at altitude, but the harsh environment precludes families from residing near the work place. Hence, workers commute between low-altitude homes and high-altitude work sites. In general, a commuting strategy is the same as the acclimatization strategy for high-altitude mountaineering, i.e., descending to low altitude when possible for rest and recovery. One goal in this strategy is to eliminate or minimize the loss of any acclimatization to hypoxia while normoxic (or less hypoxic). The benefits for different commuting schedules remain to be determined but are under active study. For example, a three-year prospective study was begun in February 1998 to characterize miners in North Chile working at 4300 to 4600 m altitude [67]. The literature oriented towards mountaineering leaves many unanswered questions about (a) the time course of acclimatization in IH; (b) the effects of different levels and duration of hypoxic exposure, i.e., the dose of IH; (c) the effects of exercise training responses to IH, and finally (d) differences in mechanisms of acclimatization to IH vs. continuous hypoxia. These issues have been addressed by measuring the time course of ventilatory and hematological changes in humans exposed to a protocol of moderate IH at rest [68,69]. The study was designed to give a lower ‘dose’ of hypoxia than previous IH studies by simulating only 3800 m altitude (PiO2 ¼ 90 Torr) for two hours per day, for 12 consecutive days, and exercise training was not part of the study [68]. The time course of change in the isocapnic HVR appeared quantitatively similar with IH and CH but on a compressed time scale with IH [69]. In IH, the isocapnic HVR (with end-tidal PCO2 held constant as inspired PO2 decreased) significantly increased to a maximum value at five days, although it subsequently decreased towards control levels by 12 days at the end of the protocol. In CH, there was a monotonic increase in isocapnic HVR during the first two weeks, similar to other studies [70]. Another difference between IH and CH was the lack of persistent hyperventilation in normoxia with IH. Despite the increase in the isocapnic HVR, ventilation and SaO2 were not significantly increased in hypoxic or normoxic conditions at any time during the IH protocol. We hypothesize that IH changed O2 sensitivity similarly to the change caused by continuous hypoxia, but it did not change normoxic ventilatory drive and arterial PCO2 set points like acclimatization to continuous hypoxia. More studies are needed comparing IH and CH to determine if fundamentally different mechanisms are involved.
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Increases in the Hypercapnic Ventilatory Response (HCVR) with Acclimatization
Some studies [70], but not all [71,72], have reported increases in the slope of hypercapnic ventilatory response (HCVR) in humans during altitude acclimatization. If one corrects for the log PCO2 relationship, the HCVR slope is not increased during acclimatization [70]. Increased HCVR slopes during acclimatization have been found in cats [30,73] and in goats [74]. As summarized by Weil [8], the mechanisms of changes in slope and intercept of the HCVR during altitude acclimatization are not entirely clear. As is the case with HVR, hyperoxia does not immediately reverse changes in the HCVR seen during acclimatization. Most of the work on the mechanisms of the increase in HCVR has focused on the carotid bodies. Acute hypoxia potentiates the carotid body neural output response to hypercapnia [75], but it is not clear whether chronic hypoxia causes further increases or not [53]. As is the case with the HVR, it remains possible that increases in central chemoreceptor sensitivity to CO2 contribute to augmented HCVR with chronic hypoxia. One possible carotid body mechanism that could increase the HCVR during chronic hypoxia is modulation of dopaminergic neurotransmission. Tatsumi, Pickett and Weil [73] found that in cats, peripheral dopaminergic blockade with domperidone increased responses to both hypoxia and hypercapnia, suggesting that decreased dopaminergic inhibition in the carotid body could explain the increase in chemosensory response in acclimatization. In these same cats, following acclimatization, a substantially diminished response to dopaminergic blockade suggests that lessening of dopaminergic activity is associated with ventilatory acclimatization. V.
High Altitude Diseases and Ventilatory Control
A. Acute Mountain Sickness (AMS)
Although AMS is more correlated with elevated heart rate than with declines in O2-hemoglobin saturation [76], it may be related to incomplete ventilatory acclimatization to altitude, since the symptoms disappear as ventilation increases (reviewed by Smith, Dempsey and Hornbein, Ref. [23]). Accordingly, it is rational to expect that the symptoms of AMS would be improved by drugs that increase ventilation or improve tissue oxygenation. Acetazolamide, a carbonic anhydrase inhibitor, improves AMS symptoms (headache, sleep disturbance, loss of appetite, malaise). Studies reporting an improvement in sleep quality and AMS symptoms with carbonic anhydrase inhibition have recently been reviewed [77]. Although it only minimally increases the HVR and HCVR [78], acetazolamide typically has its most pronounced effects in reducing periodic breathing and episodes
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of desaturation during sleep [79]. However, a recent study [80] found that arterial saturation in subjects was increased after three weeks of treatment with acetazolamide, implying an effect on ventilatory acclimatization over the longer term. The dose of acetazolamide required for benefit is 750 mg/day; 500 mg/day was not found to be effective in a 33-trial metaanalysis of drugs used for treatment of acute mountain sickness [81]. Overall, the effectiveness of acetazolamide in the treatment of the symptoms of acute mountain sickness is probably similar to that of the steroid dexamethasone [80]. The mechanism of acetazolamide’s benefit in the symptoms of acute mountain sickness may relate to improved cerebral blood flow and oxygenation that results from the carbonic acidosis it produces in brain extracellular fluids [24,81]. Almitrine bismesylate, a respiratory stimulant acting at the carotid bodies, has been investigated for its potential in speeding the process of altitude acclimatization. Although it markedly enhances ventilation and the HVR, it does not reduce periodic breathing at altitude [78]. This is very interesting because it has been proposed that an imbalance between peripheral chemoreceptor input and central drive is the basis for periodic breathing (reviewed by Weil and White, Ref. [82]). Furthermore, even though almitrine may have beneficial effects on oxygenation during wakefulness, side effects of weight loss and peripheral neuropathies may explain the lack of recent enthusiasm for the compound and its limitation for use in the treatment of chronic hypoxia [83]. Doxapram is another respiratory stimulant acting on the carotid bodies, but no information is available on the use of this drug in facilitating ventilatory acclimatization to altitude. Theophylline, a respiratory stimulant and bronchodilator, has been proposed to facilitate the respiratory acclimatization to altitude. Theophylline improves the symptoms of acute mountain sickness [76], an effect perhaps related to respiratory stimulation. Oxygenation was improved in the group receiving theophylline, but whether the effect was due to an increase in HVR or to a general increase in respiration was not delineated. Given the importance of respiratory control during sleep to the symptoms of AMS, it would be of interest to know if theophylline decreases periodic breathing or desaturations. Several studies [84,85] suggested that the extract from Ginkgo biloba might be useful for the prevention of acute mountain sickness. The mechanisms remain unknown; specifically, the respiratory effects of Gingko have not been studied. It would be interesting to determine whether the reported benefits of ginkgo correlate with enhanced blood oxygenation, ventilatory acclimatization, or improved cerebral blood flow and oxygenation. While short-acting sedative-hypnotics such as temazepam may have depressant effects on respiration at altitude [86], they may be beneficial overall because they reduce the predominant problem of periodic breathing during sleep at altitude [87]. The non-benzodiazepine sedative-hypnotic
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zolpidem has been shown to have similar benefits on sleep at altitude, without adversely influencing respiration [88]. Paradoxically, sedative drugs may improve overall gas exchange at high altitude, especially during sleep, because of their effect in reducing periodic breathing. This may result from altering the balance between peripheral and central chemoreceptor drives (see below). Several other compounds having sedative effects have been evaluated for their efficacy in treating acute mountain sickness-associated decrements in sleep quality. The mild sedative L-tryptophan does not influence acute ventilatory response at moderate altitudes [89], but its effect on AMS symptoms was not investigated. Nor is it is not known if periodic breathing or oxygen saturation are positively influenced by L-tryptophan. In contrast, alcohol, which in common with benzodiazepines produces its sedative and hypnotic effects via GABA receptors, inhibits the early stages of acute ventilatory adaptation to hypoxia at moderate altitude and is thus not recommended as a way of improving sleep quality [90]. Finally, some evidence suggests that progestational steroids such as medroxyprogesterone may reduce AMS symptoms and improve brain oxygenation [91]. B. Periodic Breathing (PB) During Sleep
Periodic breathing during sleep is a common occurrence during altitude acclimatization. It results from an imbalance in CO2- and O2-sensitive ventilatory drives. Instability arises from a low ventilatory drive during sleep and hypocapnia, with a simultaneously high ventilatory drive from hypoxia. There is a strong correlation between the HVR and PB in normal subjects at altitude [71], and the arterial chemoreceptor stimulant almitrine increases PB at altitude [78]. However, not all studies find an increase in PB with increased HVR (reviewed by Weil and White, Ref. [82]). Given the role of a high HVR in PB at altitude, it is interesting that PB decreases with acclimatization (reviewed by Weil and White, Ref. [82]) while the HVR increases (see above). However, this can be explained by the increased CO2-sensitive ventilatory drive and correction of the respiratory alkalosis during acclimatization. The increased CO2-sensitive ventilatory drive is in harmony with the elevated HVR and is able to maintain more regular breathing during sleep after acclimatization. This is similar to the ventilatory drive from wakefulness preventing PB at altitude. C. Chronic Mountain Sickness (CMS)
CMS is a poorly understood condition found in both high-altitude natives and in lowlanders living for long periods (many months to years) at high
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altitudes. Also called Monge’s disease (reviewed by Monge, Leon-Valarde and Arregui, Ref. [92]), it was first described in high-altitude residents in Bolivia and Peru. The incidence of CMS is greatest in lowlanders, and least in populations of humans genetically adapted to high-altitude life [93,94]. A primary feature of the pathophysiology of CMS appears to be a progressive loss of ventilatory drive in hypoxia, resulting in diminished ventilatory responsiveness to hypoxia, and worsening arterial oxygen saturation. Lahiri, Rozanov and Cherniak [95] reviewed the altered structure and function of the carotid bodies in chronic hypoxia. The other pathologic features of CMS very likely are caused by this progressive hypoxemia, including pulmonary hypertension, heart failure, and excessive erythrocytosis [96]. Viewed from this perspective, the disease represents a failure of respiratory control to deal with long-term environmental hypoxia, i.e., a failure of acclimatization. For unknown reasons, it affects certain individuals much more strongly than others. It would be of interest to know if individuals with pre-existing low hypoxic ventilatory responsiveness are at greater risk of developing CMS. Healthy long-term residents at high altitudes exhibit ventilation and HVR responses that are as great as recently acclimatized newcomers to altitude, indicating that the decline in ventilation in these individuals is indeed pathological, and that certain populations may be genetically advantaged with respect to maintaining ventilation during years of hypoxemia [94]. Further evidence for a fault in ventilatory drive is the correlation between the incidence of CMS and the decrease in ventilation observed with aging [92]. One prominent feature of CMS is dramatically lower arterial oxygen saturation during sleep when compared with healthy highaltitude residents. CMS patients spend a greater time in sleep-disordered breathing, with correspondingly greater periods spent desaturated, without compensatory increases in cerebral blood flow [97], indicating decreased brain oxygen delivery during a considerable portion of the night. The causes of the progressive decline in hypoxic ventilatory responses in CMS patients are not known, but the carotid bodies of patients with CMS are enlarged and structurally different than those of sea-level individuals [95,98]. There is no apparent role for endorphin-induced ventilatory decline in CMS, since naloxone does not alter the depressed ventilation of individuals with the condition [99]. There may be a role for respiratory stimulants such as medroxyprogesterone or almitrine in the hypoventilation associated with chronic mountain sickness. Medroxyprogesterone, which is effective in ameliorating both the excessive polycythemia and impaired oxygenation during sleep, may be of significant benefit. At least one study has suggested similar benefits for almitrine [100]. In general there is very little information available on the effective treatment of CMS.
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HAPE is a pathological response to hypoxia that is exacerbated by a low HVR and brisk hypoxic pulmonary vasoconstrictor response (HPVR) [101]. Susceptibility to HAPE is correlated with a low HVR in several studies [102,103], but it is important to note that not everyone with a low HVR develops HAPE. A low HVR will increase the stimulus for HAPE because alveolar PO2 will be lower for any given PiO2. Further, carotid body stimulation can reduce the HPVR so a weak HVR may lead to a stronger HPVR and increase the possibility of HAPE (101). However, these are only correlations and experiments have not established a mechanistic relationship between low ventilatory sensitivity and HAPE. References 1.
2.
3.
4. 5.
6.
7.
8.
9.
10.
Sato, M., Severinghaus, J.W., Powell, F.L., Xu, F.D. and Spellman, M.J., Jr., Augmented hypoxic ventilatory response in men at altitude, J. Appl. Physiol. 73, 101–107, 1992. Lahiri, S. and Cherniak, N.S., Cellular and molecular mechanisms of O2 sensing with special reference to the carotid body, in High Altitude: An Exploration of Human Adaptation, Hornbein, T.F. and Schoene, R.B., eds., New York, Marcel Dekker, pp. 101–130, 2001. Powell, F.L., Dwinell, M.R. and Aaron, E.A., Measuring ventilatory acclimatization to hypoxia: comparative aspects, Respir. Physiol. 122, 271–284, 2000. Powell, F.L., Milsom, W.K. and Mitchell, G.S., Time domains of the hypoxic ventilatory response, Respir. Physiol. 112, 123–134, 1998. Bisgard, G. and Neubauer, J.A., Peripheral and central effects of hypoxia, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 617–618, 1995. Eldridge, F.L. and Millhorn, D.E., Oscillation, gating, and memory in the respiratory control system, in Handbook of Physiology: The Respiratory System—Control of Breathing, Cherniack, N.S. and Widdicombe, J.G., eds. Baltimore, MD, Waverly Press, Inc, pp. 93–114, 1986. McCrimmon, D.R., Dekin, M.S. and Mitchell, G.S., Glutamate, GABA and serotonin in ventilatory control, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., New York, Marcel Dekker, pp. 151–218, 1995. Weil, J.V., Ventilatory control at high altitude, in Handbook of Physiology, Cherniak, N.S. and Widdicombe, J.G., eds., Bethesda, American Physiological Society, pp. 703–728, 1986. Smith, C.A., Dempsey, J.A. and Hornbein, T.F., Control of breathing at high altitude, in High Altitude, Hornbein, T.F. and Schoene, R.B., eds., New York, Marcel Dekker, pp. 139–173, 2002. Ohtake, P.J., Torres, J.E., Gozal, M., Graff, G.R. and Gozal D., NMDA receptors mediate peripheral chemoreceptor afferent input in the conscious rat, J. Appl. Physiol. 84, 853–861, 1998.
High Altitude 11.
12.
13. 14. 15.
16.
17. 18. 19.
20.
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25.
26.
27.
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Georgopoulos, D., Bshouty, Z., Younes, M. and Anthonisen, N.R., Hypoxic exposure and activation of the afterdischarge mechanism in conscious humans, J. Appl. Physiol. 69, 1159–1164, 1990. Badr, M.S., Skatrud, J.B. and Dempsey, J.A., Determinants of post stimulus potentiation in humans during NREM sleep, J. Appl. Physiol. 73, 1958–1971, 1992. Fregosi, R.F., Short-term potentiation of breathing in humans, J. Appl. Physiol. 71, 892–899, 1991. Wagner, P.G. and Eldridge, F.L., Development of short-term potentiation of respiration, Respir. Physiol. 83, 129–140, 1991. Hayashi, F., Coles, S.K., Bach, K.B., Mitchell, G.S. and McCrimmon, D.R., Time dependent phrenic nerve responses to carotid afferent activation: intact vs. decerebellate rats, Am. J. Physiol. 265, R811–R819, 1993. Turner, D.L. and Mitchell, G.S., Long term facilitation of ventilation following repeated hypoxic episodes in awake goats, J. Physiol. 499, 543–550, 1997. Neubauer, J.A., Melton, J.E. and Edelman, N.H., Modulation of respiration during brain hypoxia, J. Appl. Physiol. 68, 441–451, 1990. Robbins, P.A., Hypoxic ventilatory decline: site of action, J. Appl. Physiol. 79, 373–374, 1995. Bascom, D.A., Clement, I.D., Cunningham, D.A., Painter, R. and Robbins, P.A., Changes in peripheral chemoreflex sensitivity during sustained, isocapnic hypoxia, Respir. Physiol. 82, 161–176, 1990. Sato, M., Severinghaus, J.W. and Basbaum, A.I., Medullary CO2 chemoreceptor neuron acidification by c-fos immunocytochemistry, J. Appl. Physiol. 73, 96–100, 1992. Garcia, N., Hopkins, S.R., Elliott, A.R., Aaron, E.A., Weinger, M.B. and Powell, F.L., Ventilatory response to 2-h sustained hypoxia in humans, Respir. Physiol. 124, 11–22, 2000. Pedersen, M.E.F., Dorrington, K.L. and Robbins, P.A., Effects of haloperidol on ventilation during isocapnic hypoxia in humans, J. Appl. Physiol. 83, 1110–1115, 1997. Smith, C.A., Dempsey, J.A. and Hornbein, T.F., Control of breathing at high altitude, in High Altitude: An Exploration of Human Adaptation, Hornbein, T.F. and Schoene, R.B., eds., New York, Marcel Decker, pp. 139–171, 2001. Bickler, P.E., Litt, L., Banville, D.L. and Severinghaus, J.W., Effects of acetazolamide on cerebral acid-base balance, J. Appl. Physiol. 65, 422–427, 1988. Musch, T.I., Dempsey, J.A., Smith, C.A., Mitchell, G.S. and Bateman, N.T., Metabolic acids and [Hþ] regulation in brain tissue during acclimatization to chronic hypoxia, J. Appl. Physiol. 55, 1486–1495, 1983. Xu, F.D., Spellman, M.J., Jr., Mitchell, R.A. and Severinghaus, J.W., Topography of cat medullary ventral surface hypoxic acidification, J. Appl. Physiol. 73, 2631–2637, 1992. Xu, F.D., Spellman, M.J., Jr., Sato, M., Baumgartner, J.E., Circillo, S.F. and Severinghaus, J.W., Anomalous hypoxic acidification of medullary ventral surface, J. Appl. Physiol. 71, 2211–2217, 1991.
378 28. 29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42. 43. 44.
Powell and Bickler Bouverot, P., Adaptation to Altitude-Hypoxia in Vertebrates, Berlin: SpringerVerlag, 1985. Howard, L.S.G.E. and Robbins, P.A., Ventilatory response to 8 h of isocapnic and poikilocapnic hypoxia in humans, J. Appl. Physiol. 78, 1092–1097, 1995. Vizek, M., Pickett, C.K. and Weil, J.V., Increased carotid body hypoxic sensitivity during acclimitization to hypobaric hypoxia, J. Appl. Physiol. 63, 2403–2410, 1987. Forster, H.V., Dempsey, J.A., Vidruk, E.H. and Do Pico, G., Evidence of altered regulation of ventilation during exposure to hypoxia, Respir. Physiol. 20, 379–392, 1974. Powell, F.L., Huey, K.A. and Dwinell, M.R., Central nervous system mechanisms of ventilatory acclimatization to hypoxia, Respir. Physiol. 121, 223–236, 2000. Dwinell, M.R. and Powell, F.L., Chronic hypoxia enhances the phrenic nerve response to arterial chemoreceptor stimulation in anesthetized rats, J. Appl. Physiol. 87, 817–823, 1999. Schmitt, P., Soulier, V., Pequignot, J.M., Pujol, J.F. and Denavit-Saubie, M., Ventilatory acclimatization to chronic hypoxia: relationship to noradrenaline metabolism in the rat solitary complex, J. Physiol. 477, 331–337, 1994. Huey, K.A., Szewczak, J.M. and Powell, F.L., Dopaminergic mechanisms of neural plasticity in respiratory control: transgenic approaches, Respir. Physiol. Neurobiol. 135, 133–144, 2003. Huey, K.A., Low, M.J., Kelly, M.A., Juarez, R., Szewczak, J.M. and Powell, F.L., Ventilatory responses to acute and chronic hypoxia in mice: effects of dopamine D-2 receptors, J. Appl. Physiol. 89, 1142–1150, 2000. Olson, L.G. and Saunders, N.A., Effect of a dopamine antagonist on ventilation during sustained hypoxia in mice, J. Appl. Physiol. 62, 1222–1226, 1987. Dwinell, M.R., Huey, K.A. and Powell, F.L., Chronic hypoxia induces changes in the central nervous system processing of arterial chemoreceptor input, Adv. Exp. Med. Biol. 475, 477–484, 2000. Huey, K.A., Brown, I.P., Jordan, M.C. and Powell, F.L., Changes in dopamine D2-receptor modulation of the hypoxic ventilatory response with chronic hypoxia, Respir. Physiol. 123, 177–187, 2000. Mitchell, R.A., Loeschke, H.H., Massion, W.H. and Severinghaus, J.W., Respiratory responses mediated through superficial chemosensitive areas on the medulla, J. Appl. Physiol. 18, 523–533, 1963. Severinghaus, J.W., Mitchell, R.A., Richardson, B.W. and Singer, M.M., Respiratory control at high altitude suggesting active transport regulation of CSF pH, J. Appl. Physiol. 18, 1155–1166, 1963. Houston, C.S. and Riley, R.L., Respiratory and circulatory changes during acclimitization to high altitude, Am. J. Physiol. 149, 565–588, 1947. Forster, H.V. and Dempsey, J.A., Ventilatory adaptations, in Regulation of Breathing, Hornbein, T.F., ed., New York, Marcel Dekker, pp. 845–904, 1981. Crawford, R.D. and Severinghaus, J.W., CSF pH and ventilatory acclimitization to altitude, J. Appl. Physiol. 45, 275–283, 1978.
High Altitude 45. 46.
47.
48. 49. 50.
51. 52.
53.
54. 55.
56.
57.
58.
59.
60.
379
Nattie, E.E., Central chemosensitivity, sleep, and wakefulness, Resp. Physiol. 129, 257–268, 2001. Bradley, S.R., Pieribone, V.A., Wang, W., Severson, C.A., Jacobs, R.A. and Richerson, G.B., Chemosensitive serotonergic neurons are closely associated with large medullary arteries, Nat. Neurosci. 5, 401–402, 2002. Curran, L.S., Zhuang, J., Droma, T., Land, L. and Moore, L.G., Hypoxic ventilatory responses in Tibetan residents of 4400 m compared with 3658 m, Respir. Physiol. 100, 223–230, 1995. Brutsaert, T.D., Limits on inferring genetic adaptation to high altitude in Himalayan and Andean populations, High Alt. Med. Biol. 2, 211–225, 2001. Moore, L.G., Human genetic adaptation to high altitude, High Alt. Med. Biol. 2, 257–279, 2001. Rupert, J.L. and Hochachka, P.W., The evidence for hereditary factors contributing to high altitude adaptation in Andean natives: a review, High Alt. Med. Biol. 2, 235–256, 2001. Aaron, E.A. and Powell, F.L., Effect of chronic hypoxia on hypoxic ventilatory response in awake rats, J. Appl. Physiol. 74, 1635–1640, 1993. Tatsumi, K., Pickett, C.K. and Weil, J.V., Attenuated carotid body hypoxic sensitivity after prolonged hypoxic exposure, J. Appl. Physiol. 70, 748–755, 1991. Barnard, P., Andronikou, S., Pokorski, M., Smatresk, N., Mokashi, A. and Lahiri, S., Time-dependent effect of hypoxia on carotid body chemosensory function, J. Appl. Physiol. 63, 685–691, 1987. Garcia, N. and Powell, F.L., Physiological effects of intermittent hypoxia, High Alt. Med. Biol. 1, 125–136, 2000. Nagasaka, T. and Satake, T., Changes of pulmonary and cardiovascular functions in subjects confined intermittently in a low-pressure chamber for 3 consecutive days, Fed. Proc. 28, 1312–1315, 1969. Richalet, J.P., Bittel, J., Herry, J.P., Savourey, G., Le Trong, J.L., Auvert, J.F. and Janin, C., Use of a hypobaric chamber for pre-acclimatization before climbing Mount Everest, Int. J. Sports Med. 13, S216–S220, 1992. Richalet, J.P., Robach, P., Jarrot, S., Schneider, J.C., Mason, N.P., Cauchy, E., Herry, J.P., Bienvenu, A., Gardette, B. and Gortan C., Operation Everest III, in Hypoxia: Into the Next Millennium, Roach R.C., Wagner P.D. and Hackett P.H., eds., New York, Kluwer Academic/Plenum Publishing, pp. 297–317, 1999. Savourey, G., Garcia, N., Besnard, Y., Hanniquet, A.M. and Bittel, J., Pre-adaptation, adaptation and de-adaptation to high altitude in humans: cardio-ventilatory and haematological changes, Eur. J. Appl. Physiol. 73, 529–535, 1996. Benoit, H., Germain, M., Barthelemy, J.C., Denis, C., Castells, J., Dormois, D., Lacour, J.R. and Geyssant, A., Pre-acclimatization to high altitude using exercise with normobaric hypoxic gas mixtures, Int. J. Sports Med. 13, S213–S216, 1992. Katayama, K., Sato, Y., Morotome, Y., Shima, N., Ishida, K., Mori, S. and Miyamura, M., Ventilatory chemosensitive adaptations to intermittent hypoxic exposure with endurance training and detraining, J. Appl. Physiol. 86, 1805–1811, 1999.
380 61.
62.
63. 64. 65. 66.
67.
68.
69.
70.
71.
72. 73.
74.
75.
76.
Powell and Bickler Levine, B.D., Friedman, D.B., Engfred, K., Hanel, B., Kjaer, M., Clifford, P.S. and Secher, N.H., The effect of normoxic or hypobaric hypoxic endurance training on the hypoxic ventilatory response, Med. Sci. Sports Exerc. 24, 769–775, 1992. Rodriguez, F.A., Casas, H., Casas, M., Pages, T., Rama, R., Ricart, A., Ventura, J.L., Ibanez, J. and Viscor, G., Intermittent hypobaric hypoxia stimulates erythropoiesis and improves aerobic capacity, Med. Sci. Sports Exerc. 31, 264–268, 1999. West, J.B., Oxygen enrichment of room air to relieve the hypoxia of high altitude, Respir. Physiol. 99, 225–232, 1995. West, J.B., Commuting to high altitude, Adv. Exp. Med. Biol. 475, 57–64, 2000. Heath, D. and Williams, D.R., High Altitude Medicine and Physiology, Oxford, Oxford University Press, 1995. Lefrancois, R., Gautier, H., Pasquis, P. and Cavaer, A.M., Hellot, M.F. and Leroy, J., Chemoreflex ventilatory response to CO2 in man at low and high altitudes, Respir. Physiol. 14, 296–306, 1972. Richalet, J.P., Vargas, M., Jimenez, D., Moraga, F., Osorio, J., Antezana, A.M., Cauchy, E., Hudson, C., Leon, A., Cortes, G. and Brito, J., Miners exposed to intermittent high altitude hypoxia: a prospective study, Adv. Exp. Med. Biol. 474, 423 (Abstract), 1999. Garcia, N., Hopkins, S.R. and Powell, F.L., Effects of intermittent hypoxia on the isocapnic hypoxic ventilatory response and erythropoiesis in humans, Respir. Physiol. 123, 39–49, 2000. Garcia, N., Hopkins, S.R. and Powell, F.L., Intermittent versus continuous hypoxia: effects on ventilation and erythropoiesis in man, Wilderness Environ. Med. 11, 172–179, 2000. Sato, M., Severinghaus, J.W. and Bickler, P.E., Time course of augmentation and depression of hypoxic ventilatory responses at altitude, J. Appl. Physiol. 77, 313–316, 1994. White, D.P., Gleeson, K., Pickett, C.K., Rannels, A.M., Cymerman, A. and Weil, J.V., Altitude acclimatization: influence on periodic breathing and chemoresponsiveness during sleep, J. Appl. Physiol. 63, 401–412, 1987. Lahiri, S., Dynamic aspects of regulation of ventilation in man during acclimatization to high altitude, Resp. Physiol. 16, 245–258, 1972. Tatsumi, K., Pickett, C.K. and Weil, J.V., Possible role of dopamine in ventilatory acclimatization to high altitude, Resp. Physiol. 99, 63–73, 1995. Nielsen, A.M., Bisgard, G. and Vidruk, D.R., Carotid chemoreceptor activity during acute and sustained hypoxia in goats, J. Appl. Physiol. 65, 1796–1802, 1988. Hornbein, T.F., Griffo, Z.J. and Roos, A., Quantitation of chemoreceptor activity: interrelation of hypoxia and hypercapnia, J. Neurophysiol. 24, 561–568, 1961. Fischer, R., Lang, S.M., Steiner, U., Toepfer, M., Hautmann, H., Pongratz, H. and Huber, R.M., Theophylline improves acute mountain sickness, Eur. Respir. J. 15, 123–127, 2000.
High Altitude 77.
78.
79.
80.
81.
82.
83.
84.
85.
86. 87.
88.
89.
90.
91.
381
Bashir, Y., Kann, M. and Stradling, J.R., The effect of acetazolamide on hypercapnic and eucapnic/poikilocapnic hypoxic ventilatory responses in normal subjects, Pulm. Pharmacol. 3, 151–154, 1990. Hackett, P.H., Roach, R.C., Harrison, G.L., Schoene, R.B. and Mills, W.J., Jr., Respiratory stimulants and sleep periodic breathing at high altitude, Almitrine versus acetazolamide, Am. Rev. Resp. Dis. 135, 896–898, 1987. Ha, Z., Zhu, Y., Zhang, X., Cui, J., Zhang, S., Ma, Y., Wang, W. and Jian, X., The effect of rhodiola and acetazolamide on the sleep architecture and blood oxygen saturation in men living at high altitude, Zhonghua Jie He He Hu Xi Za Zhi 25, 527–530 (Chinese), 2002. Dumont, L., Mardirosoff, C. and Tramer, M.R., Efficacy and harm of pharmacological prevention of acute mountain sickness: quantitative systematic review, BMJ 321, 267–272, 2000. Bickler, P.E., Litt, L. and Severinghaus, J.W., Effects of acetazolamide on cerebrocortical NADH and blood volume, J. Appl. Physiol. 65, 428–433, 1988. Weil, J.V. and White, D.P., Sleep, in High Altitude: An Exploration of Human Adaptation, Hornbein, T.F. and Schoene, R.B., eds., New York, Marcel Dekker, pp. 707–730, 2001. Watanabe, S., Kanner, R.E., Cutillo, A.G., Menlove, R.L., Bachand, R.T., Jr., Szalkowski, M.B. and Renzetti, A.D., Long-term effect of almitrine bismesylate in patients with hypoxemic chronic obstructive pulmonary disease, Am. Rev. Respir. Dis. 140, 1269–1273, 1989. Gertsch, J.H., Seto, T.B., Mor, J. and Onopa, J., Ginkgo biloba for the prevention of severe acute mountain sickness (AMS) starting one day before rapid ascent, High Alt. Med. Biol. 3, 29–37, 2002. Roncin, J.P., Schwartz, F. and D’Arbigny, P., EGb761 in control of acute mountain sickness and vascular reactivity to cold exposure, Aviat. Space Environ. Med. 67, 445–452, 1996. Roggla, G., Moser, B. and Roggla, M., Effects of temazepam on ventilatory response at moderate altitude (Letter to editor), BMJ 320, 56, 2000. Dubowitz, G., Effect of temazepam on oxygen saturation and sleep quality at high altitude: randomized placebo controlled crossover trial, BMJ 315, 587–589, 1998. Beaumont, M., Goldenberg, F., Lejeune, D., Marotte, H., Harf, A. and Lofaso, F., Effect of zolpidem on sleep and ventilatory patterns at simulated altitude of 4,000 meters, Am. J. Resp. Crit. Care Med. 153, 1864–1869, 1996. Roggla, G., Moser, B., Wagner, A. and Roggla, M., L-Tryptophan does not influence acute ventilatory response at moderate altitude, Wein Klin. Wochenschr. 112, 634–636, 2000. Roeggla, G., Roeggla, H., Roeggla, M., Binder, M. and Laggner, A.N., Effect of alcohol on acute ventilatory adaptation to mild hypoxia at moderate altitude, Ann. Intern. Med. 122, 925–927, 1995. Imray, C.H., Barnett, N.J., Walsh, S., Clarke, T., Morgan, J., Hale, D., Hoar, H., Mole, D., Chesner, I. and Wrights, A.D., Near-infrared spectroscopy in the assessment of cerebral oxygenation at high altitude, Wilderness Environ. Med. 9, 198–203, 1998.
382
Powell and Bickler
92.
Monge, C.C., Leon-Valarde, F. and Arregui, A., Chronic mountain sickness in Andeans, in High Altitude: An Exploration of Human Adaptation, Hornbein, T.F. and Schoene, R.B., eds., New York, Marcel Dekker, pp. 815–838, 2001. Moore, L.G., Comparative human ventilatory adaptation to high altitude, Respir. Physiol. 121, 257–276, 2000. Curran, L.S., Zhuang, J. and Sun, S.F., Ventilation and hypoxic ventilatory responsiveness in Chinese-Tibetan residents at 3,658 m, J. Appl. Physiol. 83, 2098–2104, 1997. Lahiri, S., Rozanov, C. and Cherniak, N.S., Altered structure and function of the carotid body at high altitude and associated chemoreflexes, High Alt. Med. Biol. 1, 63–74, 2000. Ge, R.L. and Helun, G., Current concept of chronic mountain sickness: pulmonary hypertension-related high-altitude heart disease, Wilderness Environ. Med. 12, 190–194, 2001. Sun, S., Oliver-Pickett, C., Ping, Y., Micco, A.J., Droma, T., Zamudio, S., Zhuang, J., Huang, S.Y., McCulbugh, R.G., Cymerman, A. and Moore, L.G., Breathing and brain blood flow during sleep in patients with chronic mountain sickness, J. Appl. Physiol. 81, 611–618, 1996. Heath, D., The morbid anatomy of high altitude, Postgrad. Med. J. 55, 502–511, 1979. Sun, S.F., Huang, S.Y., Zhang, J.G., Droma, T.S., Banden, G., McCullough, R.E., Cymerman, A., Reeves, J.T. and Moore, L.G., Decreased ventilation and hypoxic ventilatory responsiveness are not reversed by naloxone in Lhasa residents with chronic mountain sickness, Am. Rev. Respir. Dis. 141, 1294–1300, 1990. Kryger, M., Glas, R., Jackson, D., McCullough, R.E., Scoggin, C., Grover, R.F. and Weil, J.V., Impaired oxygenation during sleep in excessive polycythemia of high altitude: improvement with respiratory stimulation, Sleep 1, 3–17, 1978. Schoene, R.B., Hultgren, H. and Swensen, E.R., High-altitude pulmonary edema, in High Altitude: An Exploration of Human Adaptation, Hornbein, T.F. and Schoene, R.B., eds., New York, Marcel Dekker, pp. 777–814, 2001. Selland, M.A., Stelzner, T.J., Stevens, T., Mazzeo, R.S., McCullough, R.E. and Reeves, J.T., Pulmonary function and hypoxic ventilatory response in subjects susceptible to high-altitude pulmonary edema, Chest 103, 111–116, 1993. Hackett, P.H., Roach, R.C., Schoene, R.B., Harrison, G.L. and Mills, W.J., Jr., Abnormal control of ventilation in high-altitude pulmonary edema, J. Appl. Physiol. 64, 1268–1272, 1988.
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95.
96.
97.
98. 99.
100.
101.
102.
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12 Obesity and the Control of Breathing
KHALID F. ALMOOSA
SHAHROKH JAVAHERI
University of Cincinnati College of Medicine Cincinnati VA Medical Center Cincinnati, Ohio
University of Cincinnati College of Medicine Cincinnati VA Medical Center Cincinnati, Ohio and SleepCare Diagnostics Mason, Ohio
I.
Introduction
Obesity is a common and chronic medical problem in the world today [1,2]. The health significance of obesity lies in its detrimental effects on morbidity and mortality through its direct association with several disorders [3,4]. Obesity-related health care costs in the United States amount to approximately $68 billion per year, with an additional estimated $30 billion per year spent on weight-reduction programs [5]. Obesity affects various body systems in different ways. It has numerous effects on the respiratory system, including chest wall mechanics, gas exchange, cost of breathing, and ventilatory control during both wakefulness and sleep. In this chapter, we discuss the various pathophysiological effects of obesity on pulmonary function and ventilatory control, the associated disorders of ventilatory control such as obstructive sleep apnea and the obesity hypoventilation syndrome (OHS), and the effects of obesity treatment on obstructive sleep apnea and ventilatory control.
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Overview of Obesity
A. Definition and Epidemiology
Obesity is defined as the presence of excessive body fat that causes an increase in body mass. This is reflected by an increased body-mass index (weight in kilograms divided by the square of the height in meters, BMI) [6]. A normal BMI is between 18.5 and 24.9 kg/m2, while a BMI between 25 and 29.9 kg/m2 reflects overweight, BMI greater than 30 kg/m2 defines obesity, and a BMI over 40 kg/m2 defines extreme obesity. The incidence and prevalence of obesity are increasing worldwide, particularly in industrialized countries [1,2]. Furthermore, the increase in obesity is occurring in both sexes, in all races and ages, and at all educational levels. According to Mokdad et al. (Table 12.1), the prevalence of obesity (BMI 30 kg/m2) in the United States was about 21% in 2001, while the prevalence of extreme obesity (BMI 40 kg/m2) was 2.3% [2]. Extreme obesity is highest among black women, persons who have not completed high school, and short people [7]. The prevalence of childhood obesity has also increased to 10% among 2- to 5-year olds; 15% among 6- through 11-year olds, and 16% among 12- through 19-year olds [8]. This is particularly evident among minority groups. B. Etiology and Genetics of Obesity
While the cause of obesity is incompletely understood, both genetic and environmental factors play important roles in its development [9,10]. Evidence for genetic contributions to obesity comes from both animal models [11–13] and human studies. Evidence of genetic influences in humans include highly correlated BMIs among first-degree relatives; greater BMI similarity between monozygotic vs. dizygotic twins, and significant
Table 12.1 Definitions and Prevalence of Obesity (Data from Ref. 2). Body size Normal Overweight Class I obesity Class II obesity Class III (extreme) obesity Prevalence Obesity Extreme obesity
BMI (kg/m2) 18.5–24.9 25–29.9 30–34.9 35–39.9 40 21% 2.3%
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correlation of body habitus and BMI between biologic parents and adopted children even when their environments are different [14–16]. Although obesity is a complex polygenic disorder, making it difficult to study its basis, rare genetic mutations have been identified. However, more recently, mutations in melanocortin 4 receptors, which are receptors for the anorexigenic peptide a-melanocyte, have been reported to account for 6% of childhood obesity [17]. Environmental factors have a major impact on the risk of obesity development. Western society’s affluence, easy availability of food, and the rise of fast food culture have undoubtedly all contributed to the increase in obesity over the past several decades. In addition, decreased physical activity and reduced energy expenditure associated with a sedentary life style have contributed to the development of obesity [18,19]. C. Effects of Obesity on Health
Obesity increases the risk of development of many disorders, such as adultonset diabetes mellitus, hypertension, hyperlipidemia, gall bladder disease, some cancers, and heart disease [2,20–22]. Obese subjects have twice the risk of developing heart failure as their non-obese counterparts, with risk proportional to increasing BMI (Figure 12.1) [4]. The risk of death is increased throughout the range of moderate and severe obesity for both men and women in all age groups, with risk being particularly higher for blacks than for whites (Figure 12.2) [3]. This increased risk of all-cause mortality from obesity occurs mainly through its linkage with associated diseases, particularly cardiovascular disorders [23]. Metabolic syndrome, which is linked to obesity, has been recently defined by the National Institutes of Health [24] as a constellation of two clinical and three laboratory findings. These include: (1) abdominal obesity (measured in waist circumference 4102 cm for men and 488 cm for women); (2) blood pressure (130/85 mm Hg); (3) hypertriglyceridemia (150 mg/dl); (4) low HDL (540 mg/dl for men and 550 mg/dl for women), and (5) fasting glucose 110 mg/dl. The syndrome is also associated with insulin resistance and high C-reactive protein concentration (CRP). At presentation, metabolic syndrome is defined by the presence of three or more components noted above. Metabolic syndrome is also associated with endothelial dysfunction and increased risk of cardiovascular disorders. The age-adjusted U.S. prevalence of metabolic syndrome is about 24%. As a precursor of cardiovascular diseases, detection of metabolic syndrome and its treatment may prevent associated cardiovascular pathology. Effects of obesity are not only related to the excess body fat as a percentage of total fat, but also to the distribution of fat, with the
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Cumulative incidence of heart failure (%)
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Figure 12.1 The risk of heart failure in obesity in men and women (Data from Ref. 4).
android upper-body distribution carrying a greater risk than the gynecoid lower body allocation [25]. In addition, increased neck size with fat deposition within the upper airway contributes to the presence and degree of obstructive sleep apnea–hypopnea syndrome (OSAH) [26,27].
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3.2 3.0 2.8 2.6 2.4 2.2 2.0 1.8 1.6 1.4 1.2 1.0 0.8 0.6
Cardiovascular disease Cancer All other causes
< 18 18. .5 5 – 20 20 .5 .4 – 22 21 .0 .9 – 23 23 .5 .4 – 25 24 .0 .9 – 26 26. .5 4 – 28 27. .0 9 – 30 29. .0 9 – 32 31 .0 .9 –3 4. 9 ≥3 5. 0
Relative risk of death
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Body-mass index
2.4 2.2 2.0 1.8 1.6 1.4 1.2 1.0 0.8 0.6 18 <18 .5 .5 20 −20 .5 .4 − 22 21 .0 .9 − 23 23 .5 .4 − 25 24 .0 .9 26 −26 .5 .4 28 −27 .0 .9 30 −29 .0 .9 − 32 31 .0 .9 35 −34 .0 .9 −3 9. ≥4 9 0. 0
Relative risk of death
Women
Body-mass index
Figure 12.2 The risk of death in obesity in men and women. (Data from Ref. 3).
For this reason, obesity is an important risk factor for obstructive sleep apnea. The Wisconsin Sleep Cohort Study showed that among 30- to 60-year-olds, 24% of men and 9% of women, while asleep, had an hourly rate of apnea–hypopnea (apnea–hypopnea index, AHI) score of 5 or higher [28]. In this study, an increase in BMI of one standard deviation was associated with a four-fold increase in the risk of obstructive sleep apnea.
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Sleep-related breathing disorders, in turn, are important risk factors in the development of cardiovascular disease, including systemic hypertension, coronary heart disease, heart failure, and stroke [29–31]. Furthermore, effective treatment of OSAH with nasal continuous positive airway pressure (CPAP) decreases blood pressure independently of any changes in body weight [32–34]. III.
Effects of Obesity on the Respiratory System
A. Mechanics Compliance
In obesity, total respiratory system compliance, the sum of lung compliance and chest wall compliance, is decreased [35–37], primarily due to a decrease in chest wall compliance because of accumulated fat in and around the ribs, diaphragm, and abdomen. Sharp et al. [36] confirmed this effect by measuring the respiratory system compliance while bags of buckshot were placed on the chests of supine non-obese subjects. Pelosi et al. studied the relative contributions of the lung and chest wall on the total respiratory system mechanics in 10 sedated and paralyzed morbidly obese subjects (mean BMI 49 kg/m2) and 10 normal controls (mean BMI 24 kg/m2) [38]. In the morbidly obese subjects, total respiratory system compliance was reduced by 50% (55 ml/cmH2O vs. 107 ml/cmH2O for obese and normal weight subjects, respectively). Lung compliance may also decrease in obesity in part due to increased pulmonary blood volume and microatelectasis. A reduction in the compliance of the respiratory system due to obesity may result in a pattern of lung function test abnormality referred to as a restrictive defect. The defect is characterized by decreases in functional residual capacity (FRC), expiratory reserve volume (ERV), forced vital capacity (FVC) and total lung capacity (TLC), with the forced expiratory volume in the first second (FEV1)/FVC ratio remaining normal [39,40]. These abnormalities, however, usually occur in severe obesity, with lung volumes remaining within normal range in mild to moderate obesity [39–41]. The most common abnormality seen on pulmonary function testing in obesity is a reduced ERV [40,42]. The excess thoracic and abdominal fat and cephalad movement of the diaphragm reduce FRC and ERV by the mass loading effect. This is further accentuated in the supine position, where the weight of abdominal contents pushes the diaphragm cephalad [36]. This may be the reason for orthopnea experienced by obese patients [43]. It is also important to emphasize that several studies have shown improvement in lung mechanics with weight loss [44–48]; in some of these studies, weight loss has been mild. De Lorenzo et al. measured pulmonary function in 30 obese adults (mean BMI 32 kg/m2) without underlying
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obstructive lung disease [44]. During the weight loss period, BMI decreased from 32 to 30 kg/m2 ( p 5 0.0001). With weight loss, forced expiratory flow at 50% of the volume (FEF50), FEF25–75, ERV, FEV1 (2.97 0.9 to 3.1 1 l/sec, mean SD; p 5 0.0001), and maximum voluntary ventilation (MVV) (109–122 l/min; p 5 0.0001) increased significantly. In another study, Thomas et al. measured lung volumes of 29 morbidly obese patients undergoing gastric bypass surgery [49]. Mean weight decreased from 126 to 92 kg (27% decrease), resulting in an increase in TLC by 14%, FRC by 37%, RV by 26%, FEV1 by 6%, and ERV by 54%. The FEV1/FVC ratio remained unchanged. Airways Resistance Lower airway
Obesity increases lower airway resistance. In the study by Pelosi et al., airway resistance was significantly higher (4.7 ml cmH2O l1 sec) in 10 morbidly obese subjects (mean BMI 49 kg/m2) when compared with 10 normal weight subjects (1.0 ml cmH2O l1 sec) (mean BMI 24 kg/m2) [38]. Also, Zerah et al. studied 46 patients divided into three groups based on the BMI (BMI ¼ 25–29 kg/m2, n ¼ 13; BMI ¼ 30–40 kg/m2, n ¼ 24; BMI 4 40 kg/m2, n ¼ 9) [50]. Mean airway resistance increased significantly with the level of obesity, from 3.2 cmH2O sec l1 in the first group to 5.0 cmH2O sec l1 in the third group ( p 5 0.005). The primary reason for this increased resistance is the reduction in lung volume due to obesity, since specific conductance and FEV1/FVC ratio remained normal. Upper airway
Obesity may increase fat deposition around the upper airway and in the parapharyngeal fat pads, decreasing the size of the upper airway and predisposing obese patients to obstructive sleep apnea. Animal studies involving mass loading of the neck to simulate increased neck soft tissue have demonstrated upper airway narrowing and increased resistance [51]. Increased neck circumference is a predictor of the presence and severity of sleep-related breathing disorders [52,53]; enlargement of the lateral pharyngeal walls from excess soft tissue is associated with an increased likelihood of developing sleep-related breathing disorders [26,54,55]. Horner et al. compared the neck fat deposition characteristics of six OSAH and five non-OSAH weight-matched patients and found that the OSAH patients had more fat deposition in the posterolateral pharyngeal walls surrounding the collapsible segment of the pharynx [56]. In addition, by decreasing lung volume, obesity decreases the caudal traction of the trachea on the upper airway, and may decrease upper airway size. This effect should be more pronounced in the supine position when FRC is decreased.
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The decreased respiratory system compliance and increased airway resistance in obesity requires increased neuromuscular drive for maintenance of airflow during normal breathing. Otherwise, PaCO2 should rise, particularly in the face of increased carbon dioxide production, which is also observed in obesity. Sampson et al. showed normal mean inspiratory and expiratory flow rates in 17 eucapnic massively obese subjects (mean BMI ¼ 43 kg/m2) compared with eight normal controls (mean BMI ¼ 26 kg/m2) [57]. Neuromuscular drive, as measured by mouth occlusion pressure during 0.1-sec interruption of airflow (P0.1 or P0.15), was significantly higher in the eucapnic obese patients when compared with the controls [57]. There was a significant linear correlation between P0.1, P0.1/mean inspiratory flow ratio, and percent ideal body weight. Therefore, this increased neuromuscular drive maintained normal mean inspiratory flow rate in obese subjects. The results of this study were consistent with those obtained by Burki et al., who showed that eucapnic obese subjects (n ¼ 23; mean weight ¼ 235% of ideal body weight) had increased minute ventilation and increased inspiratory neuromuscular drive when compared with non-obese controls (n ¼ 18; all were within ideal body weight range) [37]. The increase in neuromuscular drive observed in obese subjects during normal breathing, however, is not paralleled by increased maximum respiratory muscle strength. Kelly et al. measured maximum inspiratory and expiratory pressures (PIMAX and PEMAX) in 45 morbidly obese subjects (38 females, mean BMI ¼ 42 kg/m2; 7 males, mean BMI ¼ 49 kg/m2) and compared them with 25 non-obese controls (17 females, mean BMI ¼ 23 kg/m2; 8 males, mean BMI ¼ 25 kg/m2) [58]. In this study, the TLC (at which PEMAX was measured) and residual volume (at which PIMAX was measured) were similar in both groups. In both men and women, maximum pressures generated were lower (though not significantly) in obese subjects (PIMAX females: 88 cmH2O vs. 108 cmH2O controls, males: 97 cmH2O vs. 148 cmH2O controls; PEMAX females: 101 cmH2O vs. 107 cmH2O controls, males: 148 cmH2O vs. 155 cmH2O controls). Similarly, in obese subjects without hypercapnia, Lopata et al. reported that during CO2 chemostimulation, for a given increase in diaphragmatic EMG, the P0.15 was lower in obese subjects when compared with normal controls [59]. Consistent with the above reports, respiratory muscle strength increases after weight loss. Weiner et al. measured respiratory muscle strength in 21 obese subjects (mean BMI ¼ 42 kg/m2) before and six months after vertical banded gastroplasty (BMI ¼ 32 kg/m2) [48]. Consistent with Kelly et al.’s data, PIMAX and PEMAX were slightly below normal at baseline, and improved significantly with weight loss (PIMAX: 92–113 cmH2O; PEMAX: 144–166 cmH2O). We should note, however, that other studies have
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not found significantly reduced respiratory muscle strength in obesity [60]. Further, in a study involving a small number of subjects (n ¼ 16), Wadstrom et al. reported that respiratory muscle strength decreased after weight loss [47]. Ray et al. measured MVV, a reflection of muscle endurance, in 43 massively obese (mean weight 159 kg) but otherwise healthy, non-smoking, eucapnic young adults. The mean value was 123 30 l/min (mean SD, 90 22% of predicted ( p 5 0.05)) [40]. Rochester et al. studied 42 obese subjects (16 hypercapnic, mean weight ¼ 134 kg; 26 eucapnic, mean weight ¼ 138 kg) and showed that MVV was 50% predicted in the hypercapnic group and 77% predicted in the eucapnic obese group [61]. Consistent with the results of the above studies, Wadstrom et al. reported an increase in MVV40, from 94 to 107 l/min (p 5 0.001), after 23 kg weight loss in 16 obese patients (mean initial weight ¼ 125 kg; mean weight after weight loss ¼ 102 kg) [47]. Similar results were reported by Refsum et al., studying 34 women with uncomplicated obesity (mean weight ¼ 113 kg) before and one year after weight loss by gastric banding (mean weight after surgery ¼ 82 kg) [62]. MVV increased from 119 to 131 l/min after weight loss ( p 5 0.005). Rochester et al. also showed that MVV increased after weight loss, both in subjects with simple obesity and in those with OHS [61]. One study, however, found no change in MVV after weight loss [63]. Overall, from the above studies we conclude that in eucapnic obese subjects, resting neuromuscular drive is increased to compensate for increased resistance of the respiratory system. However, maximum respiratory muscle strength, P0.15 during CO2 stimulation, endurance, and diaphragmatic activity in response to CO2 stimulation are all decreased in obese subjects. Weight loss has beneficial effects on respiratory muscle function. B. O2 Consumption and CO2 Production
Obesity imposes high metabolic demands, manifested by increased O2 consumption (V_ O2 ) and CO2 production (V_ CO2 ) during both rest and exercise [63]. This is due to both excess body fat and increased energy cost of breathing because of decreased compliance and increased airway resistance [38]. Kress et al. studied the impact of morbid obesity on oxygen cost of breathing at rest [64]. Baseline V_ O2 was measured in 18 morbidly obese patients (mean BMI ¼ 53 kg/m2) and eight control subjects (mean BMI ¼ 22 kg/m2) prior to elective surgery. Baseline V_ O2 was higher in the obese group (355 ml/min) when compared with controls (221 ml/min), and the reduction in V_ O2 from spontaneous breathing to mechanical ventilation was significant in the obese patients (355–297 ml/min) but not in the control
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group (221–220 ml/min). In this study, when normalized for body surface area, there were no significant differences in V_ O2 and V_ CO2 when obese subjects were compared with normal weight subjects. These data suggest that obese individuals dedicate a larger percentage of total V_ O2 for respiratory work during quiet breathing. Pelosi et al. have reported similar results [38]. As will be discussed below, increased V_ CO2 contributes to hypercapnia associated with obesity. C. Hypoxemia, Hypercapnia, and DLCO
The increased alveolar-arterial oxygen tension difference and hypoxemia in obesity are predominantly due to V_ =Q_ mismatch [40,65,66]. Holley et al. used 133Xenon scanning to trace ventilation in obese subjects, and found that in the upright position, the distribution of tidal breathing was mainly in the upper lung zones and perfusion mainly in the lower zones [66]. Tucker et al. [67] showed a similar but more prominent effect in obese subjects in the supine position, which may account for worsening of hypoxemia in the supine position in some obese subjects. In one study, awake PaO2 was 75 mm Hg in 17 eucapnic obese men, with mean BMI ¼ 40 kg/m2 and mean age ¼ 46 years [60]. The mean PaCO2 was 39 mm Hg and the alveolar-arterial difference was 27 mm Hg. Surprisingly, in women, the mean PaO2 was significantly higher (83 mm Hg) than in men, even though the women were more obese (mean BMI ¼ 43 kg/m2) than the men. The difference in PaO2 between the two genders may have been due to difference in fat deposition, visceral vs. non-visceral, respectively, in male and female subjects. In another study, Thomas et al. reported a mean PaO2 of 79 mm Hg in 29 patients with obesity (mean weight ¼ 126 kg) [49]. In this study, PaO2 increased to 96 mm Hg after a mean of 34 kg of weight loss. Of note, 14 of these patients were either current or past smokers, and there was no indication of the presence of underlying chronic obstructive pulmonary disease (COPD) or asthma. Leech et al. reported that 70 eucapnic obese patients (53 males, 17 females) had a mean PaO2 of 72 mm Hg (range 43–99) [68]. However, some of these patients had OSAH and obstructive airways defects. In the hypercapnic patients, the mean PaCO2 was 52 mm Hg, and the mean PaO2 was 57 mm Hg. Most obese patients have normal arterial P CO2. Eucapnia is maintained, despite increased carbon dioxide production and V_ =Q_ mismatching noted above. Therefore, in eucapnic obesity, increased CO2 production and alveolar ventilation must be coupled such that arterial PCO2 remains normal. Some obese subjects, however, develop hypercapnia, and this is referred to as the OHS. Most subjects with OHS also have OSAH. The mechanisms of hypercapnia will be discussed below. In obesity, the single breath diffusion capacity (DLCO) is either normal or increased [40]. When corrected for alveolar volume, DLCO is
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usually increased [69,70], probably due to increased pulmonary capillary blood volume. DLCO decreases with weight loss [40]. D. Ventilatory Response to Hypercapnia and Hypoxemia
In order to delineate the mechanisms of hypercapnia in obesity and obstructive sleep apnea, investigators have studied hypercapnic and hypoxic ventilatory responses (HCVR, AHR). Various methods used to study HCVR and AHR are described by Ward in Chapter 4 of this volume. In general, the interpretation of ventilatory responses in subjects with obesity and pulmonary diseases is difficult, such that a decreased ventilatory response may not necessarily reflect decreased chemosensitivity. This is because, in addition to intrinsic chemosensitivity (which is partially genetically determined, see Chapter 8 by Strohl, this volume), a number of factors, such as mechanics of the respiratory system, the prevailing PO2 and PCO2, metabolic rate, medications, sleep pattern, age and gender all affect ventilatory response. Therefore, a given change in ventilatory response may not necessarily reflect changes in chemosensitivity. With these considerations, in the interpretation of studies of ventilatory responses in obese patients, several important factors need to be considered: what are the effects on ventilatory response of simple obesity (i.e., obesity without hypercapnia or concomitant OSAH), obesity with OSAH, obesity with OSAH and hypercapnia, and obesity with hypercapnia but without OSAH? Answers to these questions need particular attention with respect to appropriate control groups. For the above reasons, interpretation of the results of a number of studies, in which such information (for example, PaCO2 or polysomnography to determine the presence of sleep apnea) is not available, remains difficult. Eucapnic Obesity vs. Normal Weight
There are several studies comparing the ventilatory response to CO2 of obese subjects with normal controls, with conflicting results (Table 12.2). Lopata and Onal measured hypercapnic ventilatory responses of 34 nonobese (within 10% of IBW) and 22 obese subjects. Of the 22 obese subjects (231% of ideal weight), seven were eucapnic and without OSAH. When eucapnic obese subjects were compared with normals, obese subjects had a lower ventilatory response to CO2 (2.0 0.3 l min1 mm Hg1 vs. 3.7 0.2 l min1 mm Hg1, respectively, mean SE, p 5 0.05) [59]. Burki and Baker measured hypercapnic and hypoxic ventilatory responses in 23 eucapnic, massively obese subjects (mean age 37 years; mean body weight 149 kg, 234% of the predicted maximal ideal body weight; 16 female, 7 male) and 18 healthy normal male subjects [37]. These subjects did not undergo polysomnography, however. The overall hypercapnic ventilatory response
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Table 12.2
Comparison of Ventilatory Responses to CO2 (Slope—S, HCVR) and Hypoxemia (Slope—S, HVR) in Obesity Body size (p value)
Study groups
Author/ reference
Study group
Eucapnic obesity vs. Normal weight
Burki [37]
234% IBW N/A
Lopata [59] Kunitomoc [71]
Buyse [60]
p
Study group
Control Study group group
Control group
p
S, HVR l min1 %sat1 Study group
Control group
p
M: 18
21a
32a
50.05
44b
20b
50.01
231% IBW Within 10% IBW 50.05 M: 8 F: 186% F: 86% F: 14 IBW IBW M: 176% M: 93% IBW IBW 50.001 M: 21 M: BMI 40 N/A F: BMI 43 F: 117
M: 34
2.03
3.65
50.05
N/A
N/A
N/A
M: 15 F: 8
F: 2.2 F: 1.04 50.01 M: 1.96 M: 1.54 50.01
F: 0.15 M: 0.4
50.01 50.05
N/A
M: 3.14 N/A F: 2.37
Md: NS M: 2.46 Fd: NS F: 2.04
N/A
Md: NS Fd: 50.05
BMI 34
BMI 32
2.05
3.02
50.01
0.76
0.91
NS
BMI 31
BMI 30
M: 16 F: 1 M: 80 F: 35 M: 21 F: 34 M: 7 M: 12 F: 2
1.9
1.8
NS
N/A
N/A
N/A
M: 3.5 F: 2.1 1.68 2.41
M: 3.1 F: 1.9 2.03 1.66
NS NS NS 50.05
M: 2.8 F: 2.3 N/A N/A
M: 2.5 F: 1.7 N/A N/A
NS NS N/A N/A
M: BMI 40 F: BMI 42 Lopata [59] 191% IBW Verbraecken BMI 35 [75]
M: BMI 40 F: BMI 42 231% IBW BMI 30
N/A
M: 7 F: 16 50.05 M: 7
NS
M: 32 F: 3 0.03 M: 97 F: 7 NS M: 21 NS F: 34 50.05 M: 7 50.05 M: 12 F: 2
F: 0.9 M: 0.8
Almoosa and Javaheri
Obesity with Gold [74] OSAH vs. Obesity Sin [73] without OSAH Buyse [60]
Control group
S, HCVR l min1 mm Hg1
No. of patients
Javaheri [76] BMI 40 BMI 37 Bradley [77] 189% IBW 148% IBW
NS 23 50.05 M: 7
Lopata [59]
198% IBW Within 10% 50.05 M: 8 IBW Zwillich [81] 122 kg N/A N/A M: 10 Verbraecken BMI 33 BMI 35 NS M: 9 [75] F: 2 Garay [78] 197% IBW 177% PBW NS M: 7 Lin [80]
BMI 39.1
BMI 34.1
NS
Han [79]
BMI 27.5
BMI 29
NS
M: 5 F: 1 M: 3 F: 2
32 M: 37 F: 6 M: 7 M: 44 M: 12 F: 2 M: 5 F: 1 M: 21 F: 3 M: 3 F: 2
1.1 1.4
2.3 2.4
0.05 0.6 50.025 N/A
1.7 N/A
50.05 N/A
1.0
1.68
N/A
N/A
N/A
N/A
0.51 0.93
1.83 2.41
50.01 N/A
22e N/A
126e N/A
50.01 N/A
0.79
2.41
50.001 0.38
0.53
NS
0.46
2.59
50.05
1.49
2.97
50.05
1.32
2.18
NS
0.17
0.34
50.05
Obesity and Control of Breathing
OSAH with hypercapnia vs. OSAH without hypercapnia
IBW: Ideal body weight; PBW: Predicted body weight; BMI: Body mass index; NS: Not significant; N/A: Not available; w/h: Weight/height ratio. a Ventilation at PCO2 of 55 mm Hg. b Ventilation at saturation of 80%. c In this study, polysomnography was not performed. d Compared with reference values. e Parameter A (the smaller number indicates diminished chemosensitivity).
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was lower in obese subjects when compared with normal male controls, but the difference in ventilation only achieved significance at PCO2 of 55 mm Hg (21 vs. 32 l/min, P50.05). In contrast, the slope of hypoxic ventilatory response was significantly higher in obese subjects than normal subjects. The results on hypercapnic response were therefore similar to those of Lopata and Onal [59]. In a recent study, Buyse et al. reported the hypercapnic and hypoxic ventilatory responses in obese men and women without obstructive sleep apnea as determined by polysomnography. There were no significant differences in HCVR and AHR when comparing obese men with normal weight controls [60]. However, obese women had a significantly higher slope of both carbon dioxide and hypoxic responses when compared with nonobese women. Further, young obese women had significantly higher ventilatory responses than older obese women. Kunitomo et al. reported a similar observation [71]. These authors measured ventilatory responses in 14 obese (mean age 24 years, mean weight 108 kg) vs. eight non-obese female subjects (mean age 23, mean weight 45 kg), and eight obese (mean age 27, mean weight 122 kg) vs. 15 non-obese male subjects (mean age 28, mean weight 64 kg). In the females, the mean hypercapnic ventilatory response was significantly higher in obese than non-obese subjects (2.2 vs. 1.96 l/min, P50.01). Similarly, obese females had significantly larger hypoxic ventilatory response than non-obese women. These authors (Kunitomo et al.) reported that there were no significant differences in ventilatory responses in the male subjects with or without obesity. We should note, however, that two of the 14 females and six of the eight male subjects had mild to severe OSAH. In any case, the results of these two studies showing that obesity enhances ventilatory response in young females are consistent with the results of Burki and Baker who showed that the slope of hypoxic ventilatory response of 23 predominantly female (n ¼ 16) subjects was significantly higher than 18 controls [37]. The results of the above studies showing specific gender effects of obesity in females are also consistent with the results of a study showing a decrease in hypoxic and hypercapnic ventilatory responses with weight loss in a predominantly female group of subjects [72]. Chapman et al. measured hypercapnic and hypoxic ventilatory responses in 29 obese subjects without OSAH, predominantly female (n ¼ 26), (mean weight 123 kg, mean age 41 years) before and after gastroplasty [72]. Mean body weight fell significantly from a mean of 123 to 102 kg. The slope of hypercapnic ventilatory response decreased significantly from 1.3 to 1.1 l min1 mm Hg1 after surgery. A significant reduction in hypoxic ventilatory response was also observed. Importantly, however, when corrected for body surface area, there were no significant changes in ventilatory responses before and after weight loss.
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From the above studies, therefore, we conclude that perhaps there is a gender-specific effect of obesity on ventilatory responses. Young female patients with obesity have enhanced hypoxic and hypercapnic ventilatory responses, and these responses decrease with weight loss. Eucapnic Obesity with OSAH vs. Eucapnic Obesity without OSAH
The next question to address is the effect of OSAH on ventilatory response, and to determine if there is a gender-specific effect. Sin et al. studied 104 obese subjects with OSAH (7 women, 97 men, mean BMI 31 kg/m2; mean PCO2 37 mm Hg; mean apnea–hypopnea index, AHI ¼ 43/h) and 115 without OSAH (80 men, 35 women, mean BMI 30 kg/m2; mean PCO2 37 mm Hg; mean AHI 6/h), and found no difference in HCVR between subjects with (1.9 0.8 l min1 mm Hg1) and without OSAH (1.8 0.8 l min1 mm Hg1) [73]. However, a significant inverse relationship between age and HCVR was observed in men, and BMI was positively correlated with HCVR in women. The latter finding is consistent with results of other studies reviewed earlier [60,71], showing that obese females have enhanced hypercapnic ventilatory response. Buyse et al. studied hypercapnic and hypoxic ventilatory responses among eucapnic obese men and women with and without OSAH [60]. There were no significant differences in hypercapnic ventilatory response in 21 male obese subjects without OSA (mean age 46 years, mean BMI 40 kg/m2, mean AHI ¼ 2/h) when compared with 21 obese patients with OSAH (mean age 46 years, mean BMI 40 kg/m2, mean AHI ¼ 53/h). When females were compared, the slope of the hypoxic ventilatory response normalized for vital capacity was significantly higher in obese females with OSAH (n ¼ 34, mean age 49 years, mean BMI 42 kg/m2, mean AHI ¼ 49/h) when compared with obese females without OSAH (mean age 50 years, mean BMI 42 kg/m2, mean AHI ¼ 2/h). Results of the above studies, therefore, suggest the presence of OSAH does not significantly affect ventilatory responses, at least in male patients who maintain eucapnia. However, in female patients, OSAH may increase the hypoxic ventilatory response. In contrast to the results of the above studies, two other studies have reported that hypercapnic ventilatory response is decreased in patients with OSAH when compared with those without OSAH [59,74]. In a small but detailed study, Lopata and Onal measured hypercapnic ventilatory response, occlusion pressure, and diaphragmatic EMG in seven subjects with OSAH (weight ¼ 191% ideal body weight) and compared the results with a control group of obese subjects without OSAH (weight ¼ 231% ideal body weight) [59]. The mean values were lower for the OSAH than the non-OSAH group (Table 12.2), but the difference did not reach statistical significance.
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Gold et al. [74] studied hypercapnic and hypoxic ventilatory responses of 35 OSAH (32 men, 3 women, age 51 years; BMI kg/m2; AHI ¼ 62/h) and 17 matched non-OSAH subjects (16 men, 1 woman, age 52 years; BMI 32 kg/m2; AHI ¼ 3/h). Patients with OSAH demonstrated a lower waking ventilatory response to hypercapnia. The waking hypoxic ventilatory response, however, was not significantly different between the two groups (this study involved predominantly males). Of note, although the mean PaCO2 values for both groups were normal, patients with OSAH had a higher waking PaCO2 (40 mm Hg, range 33–45, vs. 37 mm Hg; p 5 0.001) than their non-OSAH counterparts. The authors suggested this reduction in ventilatory response and a higher waking PaCO2 in patients with OSAH was intermediate along a continuum from normal obesity to the Pickwickian syndrome (i.e., obesity with OSAH and hypercapnia). In contrast to the above studies, Verbraeken et al. reported an enhanced ventilatory response to CO2 in OSAH subjects (n ¼ 14, 12 males, age 47, BMI 35 kg/m2, AHI 53/h) compared with non-OSAH subjects (n ¼ 14, 12 males, age 45, BMI 30 kg/m2, AHI 4/h) [75]. The HCVR in the OSAH group was 2.41 0.26 l min1 mm Hg1 compared with 1.66 0.16 l min1 mm Hg1 in the non-OSAH group (mean SE, p 5 0.05). However, the OSAH subjects were significantly heavier than the non-OSAH subjects. Hypercapnic Obesity with OSAH vs. Eucapnic Obesity with OSAH
A small subset of patients with OSAH and obesity develop chronic hypercapnia. Several mechanisms may contribute to the development of hypercapnia, and several studies have reported on ventilatory response to hypercapnia and hypoxia in these subjects. Data consistently show that OSAH subjects with hypercapnia have a lower ventilatory response to both CO2 and hypoxia when compared with nonhypercapnic OSAH subjects (Table 12.2). In the largest study involving 23 hypercapnic and 32 eucapnic patients with OSAH, Javaheri et al. determined the relationship between chronic hypercapnia and the slopes of ventilatory responses to hypercapnia and hypoxia [76]. They normalized the slopes of ventilatory responses to respiratory mechanical performance of each subject, as reflected in MVV and FVC. Hypercapnic patients (PaCO2 445 mm Hg, mean ¼ 50 5 mm Hg, SD) were heavier and had more severe restrictive and obstructive defects when compared with eucapnic subjects (PaCO2 5 45 mm Hg, mean ¼ 39 3 mm Hg). They also had decreased hypoxic and hypercapnic ventilatory responses [76]. As noted earlier, mechanical impairment could have contributed to lower ventilatory responses in hypercapnic patients. However, when normalized to MVV
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(or any other spirometric test), the slope of CO2 response was still less in hypercapnic than eucapnic patients with obesity and OSAH. The results of the study by Javaheri et al. are consistent with those of Bradley et al. [77], Lopata et al. [59], Garay et al. [78], Han et al. [79], Lin et al. [80], Verbraeken et al. [75], and Zwillich et al. [81] (Table 12.2). In some of these studies, statistical significance may not have been reached because of the small numbers of patients [79–81]. These studies, therefore, collectively show that in hypercapnic patients with OSAH, both hypoxic and hypercapnic ventilatory responses are less than those in eucapnic OSAH patients. Ventilatory responses remain lower after normalization for mechanics of respiratory system [76]. The diminished ventilatory reponse, however, could be either because of chronic hypercapnia/hypoxemia and/or intrinsically decreased chemosensitivity. Chronic hypercapnia results in a rise in [HCO 3 ] in brain fluids. Consequently, during hypercapnic ventilatory response when PCO2 rises, the rise in brain [Hþ] is less than normal. Because of the confounding effects of hypercapnia on ventilatory response, the role of intrinsic chemosensitivity has been studied using normal family members of patients with and without hypercapnia. This will be discussed below. IV.
Disorders of Ventilatory Control Associated with Obesity
A. Obstructive Sleep Apnea–Hypopnea Definitions and Epidemiology
Obstructive sleep apnea–hypopnea syndrome (OSAH) is a disorder characterized by repetitive pharyngeal collapse during sleep that results in recurrent episodes of apnea or hypopnea, leading to poor sleep quality, daytime sleepiness, poor health-related quality of life, and a number of cardiovascular complications (Figure 12.3). Apnea is defined as the complete cessation of airflow for at least 10 sec, while hypopnea is a reduction in airflow, and/or thoracoabdominal excursions of 10 sec or more and associated oxygen desaturation 4%, and/or electroencephalographic evidence of arousal [82]. The number of apneas and hypopneas per hour of sleep is defined as the apnea–hypopnea index (AHI). Obstructive apneas and hypopneas are commonly observed during sleep. Young et al. showed that in the North American middle-aged working population, 24% of men and 9% of women have AHI 4 5/h [28]. In this study, 4% of men and 2% of women have an AHI 4 5/h and daytime sleepiness, which together define OSAH syndrome. Davies and Stradling analyzed 12 studies in Western populations and estimated that OSAH occurred in 1–5% of adult males [83].
400
Almoosa and Javaheri Primary event
Pathophysiological sequelae
Clinical presentation
Increased upper airway resistance
Snoring
Sleep Relaxation of pharyngeal dilator muscles Pharyngeal/airway collapse & obstructive sleep apnea/hypopnea
Witnessed apnea/hypopnea
↑ Respiratory effort & large negative PPL deflections
Nocturnal blood gas abnormalities
↑ TMP of cardiac chambers (↑ ANP), ↓ interstitial pressure Desaturationreoxygenation, Hypercapnia - hypocapnia ↑ Sympathic activity
Arousals Fragmentation of sleep Pharyngeal muscle activity restored
Resumption of airflow & correction of PO & PCO 2
2
Nocturia, ↑ lung water Pulmonary arteriolar vasoconstriction, Pulmonary HTN, Cor pulmonale, Chronic hypoventilation, Systemic HTN, LVH, CAD, Heart failure, Angina, Stroke, TIA Daytime somnolence and fatigue, wake-up unrefreshed, neuropsychiatric dysfunction
Snort, jerk
Figure 12.3 A simplified model showing sequence of events and clinical consequences in OSAH. (PPL ¼ pleural pressure; LV ¼ left ventricle; ANP ¼ atrial naturetic peptide; PO2/PCO2 ¼ partial pressure of oxygen and carbon dioxide; HTN ¼ hypertension; LVH ¼ left ventricular hypertrophy; CAD ¼ coronary artery disease; TIA ¼ transient ischemic attack; CNS ¼ central nervous system; TMP ¼ transmural pressure.)
More recently, Tishler et al. found that the 5-year incidence of mild to moderate sleep-disordered breathing is about 16% and incidence of moderately severe sleep-disordered breathing in adults is 7.5% [84]. Age, gender, BMI, waist–hip ratio, and serum cholesterol levels influenced this independently. However, with aging the effects of BMI and male gender decreased.
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The greatest risk factor for developing OSAH is obesity [28], and the risk increases with increasing weight [85]. The distribution of body fat increases the risk of developing OSAH and contributes to its severity. Millman et al. compared a group of men and women of similar BMI and waist circumference, and showed that the men had a greater severity of OSAH than the women. This was associated with increased upper body fat distribution, a larger waist–hip ratio, and a larger neck circumference in the men [86]. Pathogenesis: Control of Upper Airway Patency
Airway patency depends on the balance of forces which tend to keep the airway open versus those which tend to close it [87,88]. Patency is dictated by local anatomy and neuromuscular factors which are state-dependent (sleep vs. wakefulness) [89]. Activation of dilator muscles is an important mechanism of maintaining airway patency [87]. A number of muscles may be involved in this process, and the magnitude of the activity is determined by several mechanisms. These include input from local mechanoreceptors in the upper airway, cortex, peripheral and central chemoreceptors, and breathing centers within the brainstem [90–92]. As noted earlier, upper airway size is decreased in obese subjects when compared with normal controls. This is at least in part related to excessive fat and soft tissue surrounding the airway and in the lateral pharyngeal walls, increasing extraluminal and decreasing transmural pharyngeal pressure. Patients with sleep apnea may also have an enlarged tongue [93]. Since the pharyngeal airway is smaller in obese subjects than in normal controls [52,94], in order to maintain adequate inspiratory airflow, the upper airway muscles must be more active during wakefulness and sleep to overcome the increased resistance [95,96]. The mandatory increase in neuromuscular compensation to maintain upper airway patency in obese subjects is analogous to increased neuromuscular drive to pump muscles, necessary to provide adequate ventilation to maintain eucapnia. Sleep has a profound effect on muscles of the upper airways. In non-rapid eye movement (NREM) sleep, the reflex increase in genioglossal and tensor palatine muscle activity in response to negative pressure applied to the upper airway is reduced [97,98], and similarly, tonic activity of the tensor palatine is reduced [99]. Presumably, therefore, in patients with OSAH while asleep, neurocompensation is inadequate to maintain upper airway patency, such that in the face of negative intra-airway pressure the upper airway collapses, leading to hypopneic or apneic events during inspiration. For further discussion of this topic, see references by Richard Schwab [100] and Kingman Strohl [101].
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Another mechanism that influences upper airway patency is lung volume. Increases in lung volume help maintain upper airway patency by exerting a caudal traction on the trachea, increasing pharyngeal crosssectional area [102–104]. In contrast, the decrease in FRC associated with obesity may compromise this protective airway-dilating mechanism. Decrease in lung volume, particularly in the supine position in obese subjects, will reduce caudal traction on the upper airway, promoting pharyngeal collapsibility [104]. In this regard, waist circumference, presumably by decreasing lung volume, has been shown to correlate with sleep apnea [105]. Similarly, Tishler et al. concluded that waist–hip ratio was an independent predictor of OSAH [84].
B. OSA with Hypercapnia Prevalence of Daytime Hypercapnia
Daytime hypercapnia in OSAH occurs in a minority of patients. Most studies assessing the frequency of hypercapnia in OSAH show a prevalence of 10–20%, with higher frequencies in severely obese individuals and individuals with underlying lung disease [106]. Although diurnal hypercapnia is not common, nocturnal hypercapnia occurs regularly. It is not clear why only some patients develop diurnal hypercapnia. Nocturnal Hypercapnia
As a result of obstructive sleep apneas and hypopneas, nocturnal hypercapnia (and hypoxia) occurs intermittently. In sleep apnea– hypopnea, breathing pattern is periodic and is characterized by cessation or reduction in breathing followed by hyperpnea. Carbon dioxide accumulates during apneas and hypopneas and is cleared during hyperpnea. Therefore, during sleep, several factors contribute to how sustained and severe hypercapnia might be (Figure 12.4). These may be separated into two pathophysiological processes: those that contribute to CO2 loading and those that decrease CO2 unloading in interapneic periods. Since hypercapnia occurs when ventilation ceases (apnea) or decreases (hypopnea), with increased numbers and duration of apneas and hypopneas, the body becomes loaded with increasing amounts of carbon dioxide for longer periods. Increased metabolic CO2 production, which as noted earlier relates to body mass, tends to further increase PCO2. In other words, for a given duration of apnea, the more CO2 that is produced metabolically, the higher it raises arterial PCO2, everything else being the same. Further, as also noted earlier, obese subjects have excess CO2 production due to increased work of breathing. Therefore, during episodes of upper
Massive obesity
CO2 production
Work of breathing during episodes of upper airway obstruction and hyperpneas
CO2 loading
Intermittent Nocturnal HYPERCAPNIA
Impaired mechanics of respiratory system CO2 unloading
1. Resetting of central & peripheral chemosensitivity 2. Genetically reduced chemosensitivity
CO2 clearance Impaired parenchymal lung function Decreased hypoxic/hypercapnic ventilatory drives
Obesity and Control of Breathing
Increased number & duration of apneas & hypopneas
Chronic Diurnal HYPERCAPNIA
Hypothesized mechanisms leading to diurnal hypercapnia in OSAH.
403
Figure 12.4
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airway obstruction, increased respiratory muscle CO2 production further contributes to increased CO2 load. On the other hand, decreased CO2 clearance results in decreased CO2 unloading and more severe and sustained hypercapnia. Following obstructive apneas and hypopneas, arousals occur resulting in patency of the upper airway and resumption of ventilation. During this period, maximum CO2 clearance should occur provided that lung parenchyma, chest wall mechanics, and ventilatory response to carbon dioxide and hypoxia are normal. However, structural changes in lung parenchyma and altered mechanics that occur in various pulmonary disorders may limit CO2 clearance. Furthermore, during arousal periods, decreased hypoxic and hypercapnic ventilatory drives may decrease the magnitude of hyperpnea and compromise maximum CO2 clearance. In addition, a decrease in ventilatory response to hypoxia and hypercapnia may prolong the duration of apneas and hypopneas and contribute to excess CO2 accumulation. The balance of the above pathophysiological factors should determine the severity and duration of nocturnal hypercapnia. It is not clear, however, how intermittent nocturnal hypercapnia will eventually spill over to daytime diurnal hypercapnia. It is conceivable that those subjects in whom PCO2 remains elevated (or PO2 remains low) during most of sleep time will eventually develop diurnal hypercapnia by changing their chemosensitivity, either by resetting or diminishing chemosensitivity. However, serial measures of AHR and HCVR in untreated patients with OSAH are necessary to determine if gradual alterations in chemosensitivity occur as OSAH becomes more chronic. Such studies, however, are unethical to perform, since once OSAH is diagnosed, it should be treated. In regard to the resetting of chemosensitivity, Berthon-Jones and Sullivan have shown that after therapy with a nasal CPAP device, ventilatory response to CO2 is shifted to the left, suggesting that it had been reset [107]. In regard to the slope of hypercapnic and hypoxic ventilatory responses of patients with OSAH, however, the results of crosssectional studies have been inconsistent, with some studies concluding that OSAH decreases chemosensitivity [59,74]. Meanwhile, several studies of hypercapnic OSAH patients have shown that CO2 chemosensitivity increases with treatment of OSAH by nocturnal mechanical devices such as CPAP or bilevel positive airway pressure (BiPAP) [79,80]. Changes in (resetting or decrease in slope) hypercapnic (and hypoxic) chemosensitivity could be due to gradual and eventually sustained accumulation of HCO 3 and/or other chemical substances mediated by nocturnal hypercapnia and hypoxia [108,109]. In regard to hypoxic chemosensitivity, it has been shown that in subjects exposed to chronic hypoxemia in congenital heart disease with right to left shunt,
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chemosensitivity decreases, but is reversed after repair of the heart defect [110]. Sleep loss is another factor that could decrease chemosensitivity. White et al. measured hypoxic and hypercapnic ventilatory responses in 13 healthy men after a 24-h period of sleeplessness [111]. HCVR decreased from 2.07 0.17 (mean SE) to 1.57 0.15 l min1 mm Hg1 (p 5 0.01), and HVR decreased from 1.2 0.22 to 0.85 0.15 l min 1 1% saturation1 (p50.02). Therefore, if apneas result in cumulative sleep loss, it is also conceivable that it may diminish ventilatory response to hypercapnia and hypoxemia. Another mechanism that could mediate development of diurnal hypercapnia in patients with OSAH is diminished inherent chemosensitivity. In this regard, individuals with diminished intrinsic chemosensitivity would not be able to increase ventilation in the face of increased CO2 load, which occurs with sleep apnea and obesity. Therefore, they would be more prone to develop hypercapnia. Chemical ventilatory drives vary considerably among normal individuals, but the difference is less among members of the same family [112–114]. There is also significant correlation between the hypoxic ventilatory response of monozygotic twins when compared with non-identical twins [115,116]. The role of inherent diminished chemosensitivity mediating hypercapnia has been extensively studied in COPD. These studies suggest that inherent chemosensitivity may influence the development of hypercapnia in the face of lung disease [116–119]. It is assumed that in subjects with diminished inherent chemosensitivity that preceded the development of lung disease (e.g., COPD), CO2 retention occurs in the face of abnormal pulmonary physiology and gas exchange, resulting in increased CO2 production and diminished CO2 clearance. Similar mechanisms have been proposed for asthma [120]. It is, therefore, conceivable that in the face of obesity and increased CO2 loading, i.e., nocturnal events related to OSAH, which result in increased CO2 production, subjects with inherent decreased chemosensitivity might be prone to develop diurnal hypercapnia. Diminished chemosensitivity, in addition to decreasing the ability to clear CO2 adequately during arousals, could also contribute to prolongation of apneas and hypopneas. As a result of diminished chemosensitivity, therefore, more or less sustained nocturnal hypercapnia, increases in serum and cerebral [HCO 3 ], and eventually diurnal CO2 retention may occur. In addition, nocturnal hypoxic exposure could result in release of chemicals, which may eventually suppress ventilation [108,109,121,122]. In order to determine the role of familial chemosensitivity in mediating hypercapnia in OSAH, Javaheri et al. measured the slope of hypercapnic ventilatory response in family members of OSAH patients with and without hypercapnia [123]. They found no significant differences in the slopes of the
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family members of eucapnic and hypercapnic patients. Despite these findings, the results do not necessarily mean that subjects with inherently decreased chemosensitivity are not prone to develop hypercapnia in the face of OSAH. In any case, although most patients with OSAH are not hypercapnic, we believe that the presence of obstructive sleep apnea and of hypopnea resulting in nocturnal hypoxia and hypercapnia play a critical role in mediating daytime hypercapnia, since treatment of OSAH with either nasal CPAP or BiPAP results in normalization of PaCO2 in a number of patients (see below). Diurnal Hypercapnia
As noted above, only a small subset of patients with OSAH develops diurnal hypercapnia. This group of patients has been reported to have obesity-hypoventilation syndrome. The term hypoventilation has been erroneously used in the literature synonymously with hypercapnia. Hypoventilation, which means decreased global ventilation, is one of the mechanisms leading to hypercapnia, and systematic studies are not available to demonstrate the mechanism of hypercapnia in patients with OSAH. Physiologically, arterial PCO2 is determined by both production of CO2 and its excretion from the lungs. PaCO2 ¼ k
V_ CO2 V_ E ð1 ðVD =VT ÞÞ
where VD is dead space ventilation, VT is tidal volume, and k is the constant to make the units consistent. Therefore, for a given minute ventilation, PaCO2 will increase if the VD/VT ratio increases. This occurs when tidal volume decreases and respiratory rate increases, and is typical of patients with COPD and hypercapnia [124]. Considering the above equation, in obesity-hypoventilation syndrome, hypercapnia could be the result of one or a combination of the following factors: (a) global hypoventilation (decreased minute and alveolar ventilation, the true hypoventilators); (b) increased dead space ventilation from shallow breathing or ventilation perfusion mismatch, and (c) increased CO2 production. Studies of subjects with OSAH, both with and without hypercapnia, are necessary to determine pathophysiological mechanisms mediating hypercapnia. Although patients with obesity and hypercapnia are said to have hypoventilation syndrome, no systematic study is available to indicate this is the case. In this regard, as noted above, in patients with COPD and
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hypercapnia, global ventilation is not decreased, and hypercapnia is due to high VD/VT ratio [124]. Although studies are necessary to determine the physiological mechanisms of diurnal hypercapnia, several clinical factors have been associated with hypercapnia, some of which may be physiologically causal. Kreiger et al. studied 114 consecutive patients with OSAH, 13 of whom had hypercapnia [125]. In multiple regression analysis, the cumulative apnea duration was an independent predictor of hypercapnia. The overlap syndrome encompasses a group of patients who have both OSH and COPD. The phrase overlap syndrome was originally coined by Flenley to describe the coincident presence of OSAH with COPD [126]. This association may occur in 10–15% of patients with OSAH [77,127,128]. Not unexpectedly, these patients tend to have more gas exchange abnormalities, both hypoxemia and hypercapnia, than patients with OSAH alone [127–129]. Resta et al. compared anthropometric, pulmonary, and polysomnographic characteristics of OSAH patients with COPD [128]. For the same apnea–hypopnea index, patients with OSAH and COPD had higher PaCO2 when compared with OSAH patients without COPD. Compared with COPD patients without OSAH, they also had higher PaCO2, but also less severe obstructive impairment based on FEV1 (FEV1:FVC ratio 67% in overlap patients, 59% in COPD patients). Chaouat et al. studied 265 patients with OSAH (243 males, 22 females), and found that 11% had COPD (all males) [127]. This group had lower daytime PaO2, higher daytime PaCO2, and higher pulmonary artery pressures than patients without COPD. Leech et al. examined the relative contributions of age, gender, obesity, pulmonary function, and the severity of sleep-induced respiratory abnormalities to the development of hypercapnia in OSAH patients [68]. In 111 patients with OSAH, using stepwise logistic and multiple regression analysis, daytime PaO2, AHI, and the percent of predicted FVC were independently associated with hypercapnia. In the context of mechanisms of nocturnal hypercapnia, the unloading of CO2 will be impeded by the increased number of apneas and hypopneas and the presence of structural and mechanical alterations of the respiratory system, due to gas exchange abnormalities and the inability to maximally clear carbon dioxide (Figure 12.4). Another factor that could physiologically contribute to hypercapnia is massive obesity. In the study by Javaheri and colleagues, in OSAH patients without COPD, the hypercapnic patients were significantly more obese than non-hypercapnic subjects [52,76]. Although the actual mechanisms have not been elucidated, increased CO2 production, impairment of respiratory muscle function, and alteration in ventilatory responses could play a role.
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Effects of Treatment of Obesity and Osah on Ventilatory Control and PaCO2
Weight loss improves obesity-related morbidities and associated symptoms in the majority of people, regardless of the method used. A. Effect of Weight Loss on OSAH
Weight loss improves OSAH. Smith et al. studied two groups of obese patients with sleep apnea [130]. In the group whose weight did not change (119–120 kg), the apnea frequency did not change (66 vs. 71/h in NREM sleep, 48 vs. 48/h in REM sleep). In the 15 patients with OSAH who experienced weight loss (from 106 to 97 kg), apnea frequency decreased (55 vs. 29/h in NREM sleep, 57 vs. 38/h in REM sleep). Oxygen desaturations also decreased and sleep patterns improved with a reduction in stage I sleep and an increase in stage II sleep. Hypersomnolence also improved in 9 of 15 patients. Peiser et al. followed 15 morbidly obese patients with OSAH (mean weight 142 kg; mean AHI ¼ 82/h) referred for gastric bypass surgery [131]. Significant reduction in AHI occurred in all patients (mean AHI ¼ 15/h) two to four months after surgery (mean weight post-op 97 kg). Total sleep time, daytime hypersomnolence, and hypertension also improved. Similar findings were seen in another study of 13 morbidly obese patients (mean age 44 years; mean pre-op excess body weight 223%; mean pre-op AHI ¼ 89/h). Gastric bypass surgery results in a significant decrease in body weight (mean post-op excess body weight 150%) and in AHI (mean post-op AHI ¼ 8/h) [132]. The results of these studies therefore show that weight loss and presumably loss of fatty tissue surrounding the pharynx improves upper airway patency during sleep. With loss of tissue from the pharyngeal walls, the critical pressures necessary to keep the upper airway open decreases and therefore the number of apneas and hypopneas decrease [133–135]. Another mechanism that may contribute to upper airway patency is an increase in FRC after weight loss. As noted earlier, an increase in FRC should increase caudal traction on the upper airway [103,104]. Further, with weight loss, CO2 production should decrease. Therefore, both a reduction in the number of apneas and hypopneas, as well as decreased CO2 production all potentially contribute to convert daytime hypercapnia to eucapnia. B. Effect of Weight Loss on Ventilatory Response and PaCO2
As noted earlier, ventilatory responses decrease with weight loss. Chapman et al. measured hypoxic and hypercapnic ventilatory responses in 29 obese, mostly female patients without OSAH both before and 3–6 months after
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weight reduction by gastroplasty (weight before surgery: 123 kg, after surgery: 102 kg) [72]. The hypercapnic ventilatory response slope fell significantly after weight loss, from 2.9 2.3 to 2.2 1.1 (mean SD) l min1 mm Hg1. Hypoxic ventilatory response also decreased with weight loss. However, when hypoxic and hypercapnic responses were normalized to body surface area (BSA), the decreases were no longer significant. These findings were consistent with an earlier study of four patients by Emirgil et al. [45], suggesting that the reduction in ventilatory responses due to weight loss is secondary to a reduced metabolic rate (carbon dioxide production and oxygen consumption). Although the above studies show a decrease in HCVR with weight loss, other studies have shown that obese subjects, when compared with age matched controls, have decreased ventilatory response to CO2 [37,59,60,71]. Future studies performing serial CO2 responses in obese subjects, before and after weight loss, and in matched control subjects are necessary to clearly ascertain the effects of obesity and weight loss on HCVR and HVR. The serial studies in matched controls are necessary to control for the confounding issue of the effect of learning on serial CO2 responses. The latter element has been lacking in the design of previous studies. Since there may be gender- and age-specific effects of obesity on ventilatory responses, appropriate controls should be used. Weight loss also improves PaO2. Thomas et al. showed an increase in PaO2 with weight loss in the same group of patients discussed above, from 79 to 96 mm Hg [49]. There was a significant improvement in oxygenation, with oxygen saturation increasing from 86% to 91% in the same group of patients discussed above [62]. Weight loss may also decrease PaCO2. Rochester and Enson serially measured PaCO2 and weight in seven patients with OHS [61]. There was a correlation between the amount of weight loss and the reduction in PaCO2. Since with weight loss CO2 production decreases and apneas and hypopneas decrease, one would expect that eventually daytime hypercapnia should improve. C. Effect of OSAH Treatment with Nasal Mechanical Devices on Ventilatory Responses and PaCO2
Continuous positive airway pressure (CPAP) is the treatment of choice for OSAH [136]. CPAP increases intraluminal upper airway pressure, acting as a pneumatic splint that maintains luminal patency [137]. It also increases upper airway dimensions, reducing soft tissue edema [138]. CPAP has been effective in treating nocturnal oxygen desaturation, arrhythmias, arousals, pulmonary hypertension, and right heart failure [136]. It also decreases daytime sleepiness and improves neuropsychological function, quality of life, and presumably survival [139,140–145]. Recent studies suggest that effective use of CPAP also decreases hypertension [30,32,33].
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Rapoport et al. showed that four of eight hypercapnic obese patients (mean % ideal weight 214; PaCO2 ¼ 49 mm Hg) with OSAH returned to eucapnic levels within two weeks of treatment with nasal CPAP. Elevated serum bicarbonate levels also normalized [146]. Han et al. studied 10 OSAH patients with and without hypercapnia [79]. The mean PaCO2 was 48 mm Hg and decreased to eucapnic levels (approximately 40 mm Hg after 4–6 weeks of treatment. Hypercapnic and hypoxic responses of the hypercapnic patients increased after treatment with bilevel non-invasive ventilation. There were no significant changes in body weight (mean BMI ¼ 27.5 vs. 29 kg/m2). Lin et al. studied six hypercapnic OSAH patients (mean PaCO2 47 mm Hg; mean weight 111 kg) and 24 eucapnic OSAH patients (mean PaCO2 39 mm Hg; mean weight 96 kg) before and after two and four weeks of therapy with CPAP [80]. Nasal CPAP improved AHI in both groups (AHI before therapy ¼ 87 14/h; after therapy ¼ 8 4/h). Hypercapnic and hypoxic ventilatory responses also improved in the hypercapnic group after two weeks (HCVR ¼ 0.46 0.2 to 2.46 0.5 l min 1 kPa 1 ; AHR ¼ 1.49 0.5 to 2.81 0.9 l min1 %SaO21). The mean PaCO2 decreased from 47 to 39 mm Hg. CPAP did not have any significant effect on PaCO2 in the eucapnic group (Table 12.3). Masa et al. evaluated the effect of non-invasive ventilation (NIMV) in 22 patients with OHS (mean age 61 years; mean BMI 41 kg/m2; mean PaCO2 58 mm Hg) [147]. After 4 months of treatment with NIMV (BMI 41 kg/m2), PaCO2 decreased to 45 mm Hg ( p 5 0.001). Bilevel positive airway pressure (BiPAP) uses alternating pressures during inspiration and expiration, with inspiratory pressure being higher than expiratory pressure [148]. This permits increased ventilation, and would therefore lead to increased CO2 clearance. Waldhorn treated eight patients with hypercapnia (mean Pa CO2 60 mm Hg), three of whom had OSAH, with a nasal bilevel device [149]. After three months of therapy, mean daytime PaCO2 decreased significantly in all patients (PaCO2 decreased from 66 to 47 mm Hg). Piper et al. treated 13 obese (BMI 4 35 kg/m 2 ), hypercapnic (mean P CO 2 ¼ 62 mm Hg) OSAH patients with NIMV for 7–18 days, and showed improvements in arterial blood gas values with a rise in PaO2 from 50 to 66 mm Hg and a fall in PaCO2 values from 62 to 46 mm Hg [150]. Twelve of these patients were then able to change to CPAP therapy afterwards. In contrast to studies showing an increase in ventilatory response to CO2, Berthon-Jones et al. did not observe an increase in the slope of HCVR in 10 hypercapnic patients with OSAH (mean PCO2 ¼ 48 mm Hg; 172% of predicted weight) before and after therapy with CPAP [107]. However, they reported that the curve shifted to the left, indicating resetting of chemosensitivity.
Effect of Ventilatory Assist Devices on Daytime Hypercapnia
Body size PaCO2 Approximate (BMI) Pulmonary length of (mm Hg) function No. of Ventilatory treatment Author/ tests (weeks) Reference patients device Before After Before After p Piper [150]
13
NIMV
1–3
Lin [80]
6
CPAP
2–4
Masa [147]
22
NIMV
0.5–1
De Miguel [151]
33
CPAP
26
Han [79]
5
CPAP
4–6
FRC %pred 68 FEV1 %pred 70 FEV1/FVC ratio 84 FEV1 %pred 60 FEV1/FVC ratio 80 FEV1 %pred 54 FEV1/FVC ratio 66 FEV1 3.1 FEV1/FVC ratio 81
Slope, HCVR l min1 mm Hg1 Before After
p
Slope, HVR l min1 %sat1 Before After
p
48
–
62
46
50.05
N/A
39
38
47
38
50.05
0.46
41
41
58
45 50.001 N/A
N/A
N/A
N/A
37
34
53
45 50.001 N/A
N/A
N/A
N/A N/A N/A
28
29
48
40
1.8
NS
0.17
50.05
1.32
N/A
N/A
N/A N/A N/A
2.5 50.05 1.49
2.89 50.05
NA
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Table 12.3
N/A
0.37 N/A
CPAP: Continuous positive airway pressure; NIMV: Non-invasive mechanical ventilation; PBW: Predicted body weight; NS: Not significant.
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The effect of CPAP has also been studied in the overlap syndrome. de Miguel et al. assessed the effects of chronic CPAP therapy on lung function in 55 patients with the overlap syndrome, consisting of 33 hypercapnic (mean BMI 37 kg/m2; mean AHI ¼ 44 27/h, SE) and 22 eucapnic (mean BMI 34 kg/m2; mean AHI ¼ 29 22/h) patients [151]. After six months of therapy, there were slight but significant increases in PaO2, FEV1 (200 ml), and FVC in both groups. Furthermore, there were significant decreases in PaCO2 (53–45 mm Hg; p 5 0.001) and serum bicarbonate levels (30–26 meq/l; p 5 0.001) in the hypercapnic group. PaCO2 did not change in the eucapnic group (40 mm Hg). In summary, from the results of the above studies we conclude that treatment of OSAH with CPAP, BiPAP, and NIMV significantly improves CO2 retention and hypercapnic ventilatory response. Since the changes have occurred in relatively short periods of time after the start of treatment, this effect is most likely due to elimination of obstructive apnea and hypopnea, resulting in improvement in PaCO2 and ventilatory response. VI.
Conclusion
Obesity is a significant medical disorder that affects many people worldwide. It has numerous adverse effects on various body systems, including the respiratory system. The adverse effects of obesity on the respiratory system are apparent both during wakefulness and sleep. Obesity adversely affects the mechanics of the respiratory system, gas exchange, and, during sleep, is associated with obstructive apneas and hypopneas. Obstructive sleep apnea is a prevalent sleep-related breathing disorder that is caused by obesity. Weight loss may reverse and even eradicate these sleep-related breathing disorders. Nasal CPAP therapy has also proven to be effective in treating OSAH, and results in improved quality of life, reduction in blood pressure, and in some patients, reversal of hypercapnia.
References 1.
Kuczmarski, R.J., Flegal, K.M., Campbell, S.M. and Johnson, C.L., Increasing prevalence of overweight among US adults. The National Health and Nutrition Examination Surveys, 1960 to 1991, JAMA 272, 205–211, 1994. 2. Mokdad, A.H., Ford, E.S., Bowman, B.A., Dietz, W.H., Vinicor, F., Bales, V.S. and Marks, J.S., Prevalence of obesity, diabetes, and obesity-related health risk factors, 2001, JAMA 289, 76–79, 2003. 3. Calle, E.E., Thun, M.J., Petrelli, J.M., Rodriguez, C. and Heath, C.W., Jr., Body-mass index and mortality in a prospective cohort of U.S. adults, N. Engl. J. Med. 341, 1097–1105, 1999.
Obesity and Control of Breathing 4.
5.
6.
7.
8.
9. 10.
11.
12.
13.
14.
15.
16. 17.
18.
413
Kenchaiah, S., Evans, J.C., Levy, D., Wilson, P.W., Benjamin, E.J., Larson, M.G., Kannel, W.B. and Vasan, R.S., Obesity and the risk of heart failure, N. Engl. J. Med. 347, 305–313, 2002. Long-term pharmacotherapy in the management of obesity. National Task Force on the Prevention and Treatment of Obesity, JAMA 276, 1907–1915, 1996. Clinical Guidelines on the Identification, Evaluation, and Treatment of Overweight and Obesity in Adults—The Evidence Report. National Institutes of Health, Obes. Res. 6: Suppl 2, 51S–209S, 1998. Freedman, D.S., Khan, L.K., Serdula, M.K., Galuska, D.A. and Dietz, W.H., Trends and correlates of class 3 obesity in the United States from 1990 through 2000, JAMA 288, 1758–1761, 2002. Ogden, C.L., Flegal, K.M., Carroll, M.D. and Johnson, C.L., Prevalence and trends in overweight among US children and adolescents, 1999–2000, JAMA 288, 1728–1732, 2002. Bray, G.A., Hereditary adiposity in mice: Human lessons from the yellow and obese (OB/OB) mice, Obes. Res. 4, 91–95, 1996. Allison, D.B., Pietrobelli, A., Faith, M.S., Fontaine, K.R., Gropp, E. and Fernandez, J.R., Genetic influences on obesity, in Obesity Mechanisms and Clinical Management, Eckel, R.H., ed., Philadelphia, Lippincott Williams & Wilkins, pp. 31–74, 2003. Naggert, J.K., Fricker, L.D., Varlamov, O., Nishina, P.M., Rouille, Y., Steiner, D.F., Carroll, R.J., Paigen, B.J. and Leiter, E.H., Hyperproinsulinaemia in obese fat/fat mice associated with a carboxypeptidase E mutation which reduces enzyme activity, Nat. Genet. 10, 135–142, 1995. Zhang, Y., Proenca, R., Maffei, M., Barone, M., Leopold, L. and Friedman, J.M., Positional cloning of the mouse obese gene and its human homologue, Nature 372, 425–432, 1994. Warden, C.H., Fisler, J.S., Shoemaker, S.M., Wen, P.Z., Svenson, K.L., Pace, M.J., Lusis, A.J., Identification of four chromosomal loci determining obesity in a multifactorial mouse model, J. Clin. Investig. 95, 1545–1552, 1995. Allison, D.B., Kaprio, J., Korkeila, M., Koskenvuo, M., Neale, M.C. and Hayakawa, K., The heritability of body mass index among an international sample of monozygotic twins reared apart, Int. J. Obes. Relat. Metab. Disord. 20, 501–506, 1996. Sorensen, T.I., Holst, C., Stunkard, A.J. and Skovgaard, L.T., Correlations of body mass index of adult adoptees and their biological and adoptive relatives, Int. J. Obes. Relat. Metab. Disord. 16, 227–236, 1992. Stunkard, A.J., Foch, T.T., Hrubec, Z., A twin study of human obesity, JAMA 256, 51–54, 1986. Farooqi, I.S., Keogh, J.M., Yeo, G.S., Lank, E.J., Cheetham, T. and O’Rahilly, S., Clinical spectrum of obesity and mutations in the melanocortin 4 receptor gene, N. Engl. J. Med. 348, 1085–1095, 2003. Hu, F.B., Li, T.Y., Colditz, G.A., Willett, W.C. and Manson, J.E., Television watching and other sedentary behaviors in relation to risk of obesity and type 2 diabetes mellitus in women, JAMA 289, 1785–1791, 2003.
414 19.
20.
21. 22. 23. 24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
Almoosa and Javaheri Williamson, D.F., Madans, J., Anda, R.F., Kleinman, J.C., Kahn, H.S. and Byers, T., Recreational physical activity and ten-year weight change in a US national cohort, Int. J. Obes. Relat. Metab. Disord. 17, 279–286, 1993. Health implications of obesity, National Institutes of Health Consensus Development Conference Statement, Ann. Intern. Med. 103, 147–151, 1985. Allison, D.B. and Saunders, S.E., Obesity in North America. An overview, Med. Clin. North Am. 84, 305–332, v, 2000. Foster, G.D., Principles and practices in the management of obesity, Am. J. Respir. Crit. Care Med. 168, 274–280, 2003. Manson, J.E., Stampfer, M.J., Hennekens, C.H. and Willett, W.C., Body weight and longevity. A reassessment, JAMA 257, 353–358, 1987. Third Report of the National Cholesterol Education Program (NCEP) Expert Panel on Detection, Evaluation, and Treatment of High Blood Cholesterol in Adults (Adult Treatment Panel III) Final Report, Circulation 106, 3143–3421, 2002. Krotkiewski, M., Bjorntorp, P., Sjostrom, L. and Smith, U., Impact of obesity on metabolism in men and women. Importance of regional adipose tissue distribution, J. Clin. Investig. 72, 1150–1162, 1983. Mortimore, I.L., Marshall, I., Wraith, P.K., Sellar, R.J. and Douglas, N.J., Neck and total body fat deposition in non-obese and obese patients with sleep apnea compared with that in control subjects, Am. J. Respir. Crit. Care Med. 157, 280–283, 1998. Schafer, H., Pauleit, D., Sudhop, T., Gouni-Berthold, I., Ewig, S. and Berthold, H.K., Body fat distribution, serum leptin, and cardiovascular risk factors in men with obstructive sleep apnea, Chest 122, 829–839, 2002. Young, T., Palta, M., Dempsey, J., Skatrud, J., Weber, S. and Badr, S., The occurrence of sleep-disordered breathing among middle-aged adults, N. Engl. J. Med. 328, 1230–1235, 1993. Nieto, F.J., Young, T.B., Lind, B.K., Shahar, E., Samet, J.M., Redline, S., D’Agostino, R.B., Newman, A.B., Lebowitz, M.D. and Pickering, T.G., Association of sleep-disordered breathing, sleep apnea, and hypertension in a large community-based study. Sleep Heart Health Study, JAMA 283, 1829– 1836, 2000. Peppard, P.E., Young, T., Palta, M. and Skatrud, J., Prospective study of the association between sleep-disordered breathing and hypertension, N. Engl. J. Med. 342, 1378–1384, 2000. Shahar, E., Whitney, C.W., Redline, S., Lee, E.T., Newman, A.B., Javier Nieto, F., O’Connor, G.T., Boland, L.L., Schwartz, J.E. and Samet, J.M., Sleepdisordered breathing and cardiovascular disease: Cross-sectional results of the Sleep Heart Health Study, Am. J. Respir. Crit. Care Med. 163, 19–25, 2001. Becker, H.F., Jerrentrup, A., Ploch, T., Grote, L., Penzel, T., Sullivan, C.E. and Peter, J.H., Effect of nasal continuous positive airway pressure treatment on blood pressure in patients with obstructive sleep apnea, Circulation 107, 68–73, 2003. Faccenda, J.F., Mackay, T.W., Boon, N.A. and Douglas, N.J., Randomized placebo-controlled trial of continuous positive airway pressure on blood
Obesity and Control of Breathing
34.
35. 36. 37. 38.
39. 40. 41.
42. 43.
44.
45.
46.
47.
48.
49.
415
pressure in the sleep apnea–hypopnea syndrome, Am. J. Respir. Crit. Care Med. 163, 344–348, 2001. Pepperell, J.C., Ramdassingh-Dow, S., Crosthwaite, N., Mullins, R., Jenkinson, C., Stradling, J.R. and Davies, R.J., Ambulatory blood pressure after therapeutic and subtherapeutic nasal continuous positive airway pressure for obstructive sleep apnoea: A randomised parallel trial, Lancet 359, 204–210, 2002. Naimark, A. and Cherniack, R.M., Compliance of the respiratory system and its components in health and obesity, J. Clin. Investig. 15, 377–382, 1960. Sharp, J.T., Henry, J.P. and Seanhy, S.K., The total work of breathing in normal and obese men, J. Clin. Investig. 43, 728–739, 1964. Burki, N.K. and Baker, R.W., Ventilatory regulation in eucapnic morbid obesity, Am. Rev. Respir. Dis. 129, 538–543, 1984. Pelosi, P., Croci, M., Ravagnan, I., Vicardi, P. and Gattinoni, L., Total respiratory system, lung, and chest wall mechanics in sedated-paralyzed postoperative morbidly obese patients, Chest 109, 144–151, 1996. Lazarus, R., Sparrow, D. and Weiss, S.T., Effects of obesity and fat distribution on ventilatory function: The normative aging study, Chest 111, 891–898, 1997. Ray, C.S., Sue, D.Y., Bray, G., Hansen, J.E. and Wasserman, K., Effects of obesity on respiratory function, Am. Rev. Respir. Dis. 128, 501–506, 1983. Collins, L.C., Hoberty, P.D., Walker, J.F., Fletcher, E.C. and Peiris, A.N., The effect of body fat distribution on pulmonary function tests, Chest 107, 1298–1302, 1995. Bedell, G.N., Wilson, W.R. and Seebohn, P., Pulmonary function in obese persons, J. Clin. Investig. 37, 1049, 1958. Ferretti, A., Giampiccolo, P., Cavalli, A., Milic-Emili, J. and Tantucci, C., Expiratory flow limitation and orthopnea in massively obese subjects, Chest 119, 1401–1408, 2001. De Lorenzo, A., Maiolo, C., Mohamed, E.I., Andreoli, A., Petrone-De Luca, P. and Rossi, P., Body composition analysis and changes in airways function in obese adults after hypocaloric diet, Chest 119, 1409–1415, 2001. Emirgil, C. and Sobol, B.J., The effects of weight reduction on pulmonary function and the sensitivity of the respiratory center in obesity, Am. Rev. Respir. Dis. 108, 831–842, 1973. Stenius-Aarniala, B., Poussa, T., Kvarnstrom, J., Gronlund, E.L., Ylikahri, M. and Mustajoki, P., Immediate and long term effects of weight reduction in obese people with asthma: Randomised controlled study, BMJ 320, 827–832, 2000. Wadstrom, C., Muller-Suur, R. and Backman, L., Influence of excessive weight loss on respiratory function. A study of obese patients following gastroplasty, Eur. J. Surg. 157, 341–346, 1991. Weiner, P., Waizman, J., Weiner, M., Rabner, M., Magadle, R. and Zamir, D., Influence of excessive weight loss after gastroplasty for morbid obesity on respiratory muscle performance, Thorax 53, 39–42, 1998. Thomas, P.S., Cowen, E.R., Hulands, G. and Milledge, J.S., Respiratory function in the morbidly obese before and after weight loss, Thorax 44, 382– 386, 1989.
416 50. 51. 52.
53.
54.
55.
56.
57. 58. 59. 60.
61.
62.
63. 64.
65.
Almoosa and Javaheri Zerah, F., Harf, A., Perlemuter, L., Lorino, H., Lorino, A.M. and Atlan, G., Effects of obesity on respiratory resistance, Chest 103, 1470–1476, 1993. Koenig, J.S. and Thach, B.T., Effects of mass loading on the upper airway, J. Appl. Physiol. 64, 2294–2299, 1988. Schwab, R.J., Gefter, W.B., Hoffman, E.A., Gupta, K.B. and Pack, A.I., Dynamic upper airway imaging during awake respiration in normal subjects and patients with sleep disordered breathing, Am. Rev. Respir. Dis. 148, 1385– 1400, 1993. Sforza, E., Petiau, C., Weiss, T., Thibault, A. and Krieger, J., Pharyngeal critical pressure in patients with obstructive sleep apnea syndrome. Clinical implications, Am. J. Respir. Crit. Care Med. 159, 149–157, 1999. Schellenberg, J.B., Maislin, G. and Schwab, R.J., Physical findings and the risk for obstructive sleep apnea. The importance of oropharyngeal structures, Am. J. Respir. Crit. Care Med. 162, 740–748, 2000. Schwab, R.J., Gupta, K.B., Gefter, W.B., Metzger, L.J., Hoffman, E.A. and Pack, A.I., Upper airway and soft tissue anatomy in normal subjects and patients with sleep-disordered breathing. Significance of the lateral pharyngeal walls, Am. J. Respir. Crit. Care Med. 152, 1673–1689, 1995. Horner, R.L., Mohiaddin, R.H., Lowell, D.G., Shea, S.A., Burman, E.D., Longmore, D.B. and Guz, A., Sites and sizes of fat deposits around the pharynx in obese patients with obstructive sleep apnoea and weight matched controls, Eur. Respir. J. 2, 613–622, 1989. Sampson, M.G. and Grassino, A.E., Load compensation in obese patients during quiet tidal breathing, J. Appl. Physiol. 55, 1269–1276, 1983. Kelly, T.M., Jensen, R.L., Elliott, C.G. and Crapo, R.O., Maximum respiratory pressures in morbidly obese subjects, Respiration 54, 73–77, 1988. Lopata, M. and Onal, E., Mass loading, sleep apnea, and the pathogenesis of obesity hypoventilation, Am. Rev. Respir. Dis. 126, 640–645, 1982. Buyse, B., Markous, N., Cauberghs, M., Van Klaveren, R., Muls, E. and Demedts, M., Effect of obesity and/or sleep apnea on chemosensitivity: Differences between men and women, Respir. Physiol. Neurobiol. 134, 13–22, 2003. Rochester, D.F. and Enson, Y., Current concepts in the pathogenesis of the obesity-hypoventilation syndrome. Mechanical and circulatory factors, Am. J. Med. 57, 402–420, 1974. Refsum, H.E., Holter, P.H., Lovig, T., Haffner, J.F. and Stadaas, J.O., Pulmonary function and energy expenditure after marked weight loss in obese women: Observations before and one year after gastric banding, Int. J. Obes. 14, 175–183, 1990. Gilbert, R., Sipple, J.H. and Auchincloss, J.H., Respiratory control and work of breathing in obese subjects, J. Appl. Physiol. 16, 21–26, 1961. Kress, J.P., Pohlman, A.S., Alverdy, J. and Hall, J.B., The impact of morbid _O obesity on oxygen cost of breathing ðV Þ at rest, Am. J. Respir. Crit. Care 2RESP Med. 160, 883–886, 1999. Douglas, F.G. and Chong, P.Y., Influence of obesity on peripheral airways patency, J. Appl. Physiol. 33, 559–563, 1972.
Obesity and Control of Breathing 66.
67.
68. 69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
417
Holley, H.S., Milic-Emili, J., Becklake, M.R. and Bates, D.V., Regional distribution of pulmonary ventilation and perfusion in obesity, J. Clin. Investig. 46, 475, 1967. Tucker, D.H. and Sieker, H.O., The effects of change in body position on lung volumes and intrapulmonary gas mixing in patients with obesity, heart failure, and emphysema, J. Clin. Investig. 39, 787–789, 1960. Leech, J.A., Onal, E., Baer, P. and Lopata, M., Determinants of hypercapnia in occlusive sleep apnea syndrome, Chest 92, 807–813, 1987. Baylor, P. and Goebel, P., Clinical correlates of an elevated diffusing capacity for carbon monoxide corrected for alveolar volume, Am. J. Med. Sci. 311, 266–271, 1996. Collard, P., Wilputte, J.Y., Aubert, G., Rodenstein, D.O. and Frans, A., The single-breath diffusing capacity for carbon monoxide in obstructive sleep apnea and obesity, Chest 110, 1189–1193, 1996. Kunitomo, F., Kimura, H., Tatsumi, K., Kuriyama, T., Watanabe, S. and Honda, Y., Sex differences in awake ventilatory drive and abnormal breathing during sleep in eucapnic obesity, Chest 93, 968–976, 1988. Chapman, K.R., Himal, H.S. and Rebuck, A.S., Ventilatory responses to hypercapnia and hypoxia in patients with eucapnic morbid obesity before and after weight loss, Clin. Sci. (Lond.) 78, 541–545, 1990. Sin, D.D., Jones, R.L. and Man, G.C., Hypercapnic ventilatory response in patients with and without obstructive sleep apnea: Do age, gender, obesity, and daytime PaCO2 matter? Chest 117, 454–459, 2000. Gold, A.R., Schwartz, A.R., Wise, R.A. and Smith, P.L., Pulmonary function and respiratory chemosensitivity in moderately obese patients with sleep apnea, Chest 103, 1325–1329, 1993. Verbraecken, J., De Backer, W., Willemen, M., De Cock, W., Wittesaele, W. and Van de Heyning, P., Chronic CO2 drive in patients with obstructive sleep apnea and effect of CPAP, Respir. Physiol. 101, 279–287, 1995. Javaheri, S., Colangelo, G., Lacey, W. and Gartside, P.S., Chronic hypercapnia in obstructive sleep apnea–hypopnea syndrome, Sleep 17, 416–423, 1994. Bradley, T.D., Rutherford, R. and Lue, F., Role of diffuse airway obstruction in the hypercapnia of obstructive sleep apnea, Am. Rev. Respir. Dis. 134, 920–924, 1986. Garay, S.M., Rapoport, D., Sorkin, B., Epstein, H., Feinberg, I. and Goldring, R.M., Regulation of ventilation in the obstructive sleep apnea syndrome, Am. Rev. Respir. Dis. 124, 451–457, 1981. Han, F., Chen, E., Wei, H., He, Q., Ding, D. and Strohl, K.P., Treatment effects on carbon dioxide retention in patients with obstructive sleep apnea– hypopnea syndrome, Chest 119, 1814–1819, 2001. Lin, C.C., Effect of nasal CPAP on ventilatory drive in normocapnic and hypercapnic patients with obstructive sleep apnoea syndrome, Eur. Respir. J. 7, 2005–2010, 1994. Zwillich, C.W., Sutton, F.D., Pierson, D.J., Greagh, E.M. and Weil, J.V., Decreased hypoxic ventilatory drive in the obesity-hypoventilation syndrome, Am. J. Med. 59, 343–348, 1975.
418 82.
83. 84.
85.
86.
87.
88. 89. 90.
91.
92.
93.
94. 95.
96.
Almoosa and Javaheri Sleep-related breathing disorders in adults: Recommendations for syndrome definition and measurement techniques in clinical research. The Report of an American Academy of Sleep Medicine Task Force, Sleep 22, 667–689 1999. Davies, R.J. and Stradling, J.R., The epidemiology of sleep apnoea, Thorax 51: Suppl 2, S65–S70, 1996. Tishler, P.V., Larkin, E.K., Schluchter, M.D. and Redline, S., Incidence of sleep-disordered breathing in an urban adult population: The relative importance of risk factors in the development of sleep-disordered breathing, JAMA 289, 2230–2237, 2003. Vgontzas, A.N., Tan, T.L., Bixler, E.O., Martin, L.F., Shubert, D. and Kales, A., Sleep apnea and sleep disruption in obese patients, Arch. Intern. Med. 154, 1705–1711, 1994. Millman, R.P., Carlisle, C.C., McGarvey, S.T., Eveloff, S.E. and Levinson, P.D., Body fat distribution and sleep apnea severity in women, Chest 107, 362– 366, 1995. Kuna, S. and Remmers, J.E., Anatomy and physiology of upper airway obstruction, in Principles and Practice of Sleep Medicine, Kryger, M., Roth, T. and Dement, W., eds., Philadelphia, W.B Saunders Company, pp. 840–858, 2000. Badr, M.S., Effect of ventilatory drive on upper airway patency in humans during NREM sleep, Respir. Physiol. 103, 1–10, 1996. Remmers, J.E., Degroot, W.J., Sauerland, E.K. and Anch, A.M., Pathogenesis of upper airway occlusion during sleep, J. Appl. Physiol. 44, 931–938, 1978. Malhotra, A., Pillar, G., Fogel, R.B., Edwards, J.K., Ayas, N., Akahoshi, T., Hess, D. and White, D.P., Pharyngeal pressure and flow effects on genioglossus activation in normal subjects, Am. J. Respir. Crit. Care Med. 165, 71–77, 2002. Malhotra, A., Fogel, R.B., Edwards, J.K., Shea, S.A. and White, D.P., Local mechanisms drive genioglossus activation in obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 161, 1746–1749, 2000. Malhotra, A., Pillar, G., Fogel, R.B., Beauregard, J., Edwards, J.K., Slamowitz, D.I., Shea, S.A. and White, D.P., Genioglossal but not palatal muscle activity relates closely to pharyngeal pressure, Am. J. Respir. Crit. Care Med. 162, 1058–1062, 2000. Ryan, C.F., Lowe, A.A., Li, D. and Fleetham, J.A., Three-dimensional upper airway computed tomography in obstructive sleep apnea. A prospective study in patients treated by uvulopalatopharyngoplasty, Am. Rev. Respir. Dis. 144, 428–432, 1991. Fleetham, J.A., Upper airway imaging in relation to obstructive sleep apnea, Clin. Chest Med. 13, 399–416, 1992. Mezzanotte, W.S., Tangel, D.J. and White, D.P., Waking genioglossal electromyogram in sleep apnea patients versus normal controls (a neuromuscular compensatory mechanism), J. Clin. Investig. 89, 1571–1579, 1992. Horner, R.L., Innes, J.A., Murphy, K. and Guz, A., Evidence for reflex upper airway dilator muscle activation by sudden negative airway pressure in man, J. Physiol. 436, 15–29, 1991.
Obesity and Control of Breathing 97.
98.
99.
100. 101. 102.
103. 104. 105.
106.
107.
108. 109.
110.
111.
112.
113.
419
Wheatley, J.R., Mezzanotte, W.S., Tangel, D.J. and White, D.P., Influence of sleep on genioglossus muscle activation by negative pressure in normal men, Am. Rev. Respir. Dis. 148, 597–605, 1993. Wheatley, J.R., Tangel, D.J., Mezzanotte, W.S. and White, D.P., Influence of sleep on response to negative airway pressure of tensor palatini muscle and retropalatal airway, J. Appl. Physiol. 75, 2117–2124, 1993. Tangel, D.J., Mezzanotte, W.S. and White, D.P., Influence of sleep on tensor palatini EMG and upper airway resistance in normal men, J. Appl. Physiol. 70, 2574–2581, 1991. Schwab, R.J., Pro: Sleep apnea is an anatomic disorder, Am. J. Respir. Crit. Care Med. 168, 270–271, 2003. Strohl, K.P., Con: Sleep apnea is not an anatomic disorder, Am. J. Respir. Crit. Care Med. 168, 271–272, 2003. Begle, R.L., Badr, S., Skatrud, J.B. and Dempsey, J.A., Effect of lung inflation on pulmonary resistance during NREM sleep, Am. Rev. Respir. Dis. 141, 854–860, 1990. Van de Graaff, W.B., Thoracic influence on upper airway patency, J. Appl. Physiol. 65, 2124–2131, 1988. Van de Graaff, W.B., Thoracic traction on the trachea: Mechanisms and magnitude, J. Appl. Physiol. 70, 1328–1336, 1991. Grunstein, R., Wilcox, I., Yang, T.S., Gould, Y. and Hedner, J., Snoring and sleep apnoea in men: Association with central obesity and hypertension, Int. J. Obes. Relat. Metab. Disord. 17, 533–540, 1993. Weitzenblum, E., Chaouat, A., Kessler, R., Oswald, M., Apprill, M. and Krieger, J., Daytime hypoventilation in obstructive sleep apnoea syndrome, Sleep Med. Rev. 3, 79–93, 1999. Berthon-Jones, M. and Sullivan, C.E., Time course of change in ventilatory response to CO2 with long-term CPAP therapy for obstructive sleep apnea, Am. Rev. Respir. Dis. 135, 144–147, 1987. Javaheri, S. and Teppema, L.J., Ventral medullary extracellular fluid pH and PCO2 during hypoxemia, J. Appl. Physiol. 63, 1567–1571, 1987. Javaheri, S., Teppema, L.J. and Evers, J.A., Effects of aminophylline on hypoxemia-induced ventilatory depression in the cat, J. Appl. Physiol. 64, 1837–1843, 1988. Edelman, N.H., Lahiri, S., Braudo, L., Cherniack, N.S. and Fishman, A.P., The blunted ventilatory response to hypoxia in cyanotic congenital heart disease, N. Engl. J. Med. 282, 405–411, 1970. White, D.P., Douglas, N.J., Pickett, C.K., Zwillich, C.W. and Weil, J.V., Sleep deprivation and the control of ventilation, Am. Rev. Respir. Dis. 128, 984–986, 1983. Hirshman, C.A., McCullough, R.E. and Weil, J.V., Normal values for hypoxic and hypercapnic ventilatory drives in man, J. Appl. Physiol. 38, 1095–1098, 1975. Saunders, N.A., Leeder, S.R. and Rebuck, A.S., Ventilatory response to carbon dioxide in young athletes: A family study, Am. Rev. Respir. Dis. 113, 497–502, 1976.
420
Almoosa and Javaheri
114. Scoggin, C.H., Doekel, R.D., Kryger, M.H., Zwillich, C.W. and Weil, J.V., Familial aspects of decreased hypoxic drive in endurance athletes, J. Appl. Physiol. 44, 464–468, 1978. 115. Collins, D.D., Scoggin, C.H., Zwillich, C.W. and Weil, J.V., Hereditary aspects of decreased hypoxic response, J. Clin. Investig. 62, 105–110, 1978. 116. Kawakami, Y., Yoshikawa, T., Shida, A., Asanuma, Y. and Murao, M., Control of breathing in young twins, J. Appl. Physiol. 52, 537–542, 1982. 117. Moore, G.C., Zwillich, C.W., Battaglia, J.D., Cotton, E.K. and Weil, J.V., Respiratory failure associated with familial depression of ventilatory response to hypoxia and hypercapnia, N. Engl. J. Med. 295, 861–865, 1976. 118. Mountain, R., Zwillich, C. and Weil, J., Hypoventilation in obstructive lung disease. The role of familial factors, N. Engl. J. Med. 298, 521–525, 1978. 119. Hudgel, D.W. and Devadatta, P., Decrease in functional residual capacity during sleep in normal humans, J. Appl. Physiol. 57, 1319–1322, 1984. 120. Fleetham, J.A., Arnup, M.E. and Anthonisen, N.R., Familial aspects of ventilatory control in patients with chronic obstructive pulmonary disease, Am. Rev. Respir. Dis. 129, 3–7, 1984. 121. Eldridge, F.L., Millhorn, D.E. and Kiley, J.P., Antagonism by theophylline of respiratory inhibition induced by adenosine, J. Appl. Physiol. 59, 1428–1433, 1985. 122. Millhorn, D.E., Eldridge, F.L., Kiley, J.P. and Waldrop, T.G., Prolonged inhibition of respiration following acute hypoxia in glomectomized cats, Respir. Physiol. 57, 331–340, 1984. 123. Javaheri, S., Colangelo, G., Corser, B. and Zahedpour, M., Familial respiratory chemosensitivity does not predict hypercapnia of patients with sleep apnea–hypopnea syndrome, Am. Rev. Respir. Dis. 145, 837–840, 1992. 124. Javaheri, S., Blum, J. and Kazemi, H., Pattern of breathing and carbon dioxide retention in chronic obstructive lung disease, Am. J. Med. 71, 228–234, 1981. 125. Krieger, J., Sforza, E., Apprill, M., Lampert, E., Weitzenblum, E. and Ratomaharo, J., Hypercapnia in obstructive sleep apnea patients, Chest 96, 729–737, 1989. 126. Flenley, D.C., Sleep in chronic obstructive lung disease, Clin. Chest Med. 6, 651–661, 1985. 127. Chaouat, A., Weitzenblum, E., Krieger, J., Ifoundza, T., Oswald, M. and Kessler, R., Association of chronic obstructive pulmonary disease and sleep apnea syndrome, Am. J. Respir. Crit. Care Med. 151, 82–86, 1995. 128. Resta, O., Foschino Barbaro, M.P., Brindicci, C., Nocerino, M.C., Caratozzolo, G. and Carbonara, M., Hypercapnia in overlap syndrome: Possible determinant factors, Sleep Breath 6, 11–18, 2002. 129. Fletcher, E.C., Schaaf, J.W., Miller, J. and Fletcher, J.G., Long-term cardiopulmonary sequelae in patients with sleep apnea and chronic lung disease, Am. Rev. Respir. Dis. 135, 525–533, 1987. 130. Smith, P.L., Gold, A.R., Meyers, D.A., Haponik, E.F. and Bleecker, E.R., Weight loss in mildly to moderately obese patients with obstructive sleep apnea, Ann. Intern. Med. 103, 850–855, 1985.
Obesity and Control of Breathing
421
131. Peiser, J., Lavie, P., Ovnat, A. and Charuzi, I., Sleep apnea syndrome in the morbidly obese as an indication for weight reduction surgery, Ann. Surg. 199, 112–115, 1984. 132. Charuzi, I., Ovnat, A., Peiser, J., Saltz, H., Weitzman, S. and Lavie, P., The effect of surgical weight reduction on sleep quality in obesity-related sleep apnea syndrome, Surgery 97, 535–538, 1985. 133. Rubinstein, I., Colapinto, N., Rotstein, L.E., Brown, I.G. and Hoffstein, V., Improvement in upper airway function after weight loss in patients with obstructive sleep apnea, Am. Rev. Respir. Dis. 138, 1192–1195, 1988. 134. Schwartz, A.R., Gold, A.R., Schubert, N., Stryzak, A., Wise, R.A., Permutt, S. and Smith, P.L., Effect of weight loss on upper airway collapsibility in obstructive sleep apnea, Am. Rev. Respir. Dis. 144, 494–498, 1991. 135. Suratt, P.M., McTier, R.F., Findley, L.J., Pohl, S.L. and Wilhoit, S.C., Changes in breathing and the pharynx after weight loss in obstructive sleep apnea, Chest 92, 631–637, 1987. 136. Indications and standards for use of nasal continuous positive airway pressure (CPAP) in sleep apnea syndromes. American Thoracic Society. Official statement adopted March 1944, Am. J. Respir. Crit. Care Med. 150, 1738–1745, 1994. 137. Sullivan, C.E., Issa, F.G., Berthon-Jones, M. and Eves, L., Reversal of obstructive sleep apnoea by continuous positive airway pressure applied through the nares, Lancet 1, 862–865, 1981. 138. Ryan, C.F., Lowe, A.A., Li, D. and Fleetham, J.A., Magnetic resonance imaging of the upper airway in obstructive sleep apnea before and after chronic nasal continuous positive airway pressure therapy, Am. Rev. Respir. Dis. 144, 939–944, 1991. 139. Derderian, S.S., Bridenbaugh, R.H. and Rajagopal, K.R., Neuropsychologic symptoms in obstructive sleep apnea improve after treatment with nasal continuous positive airway pressure, Chest 94, 1023–1027, 1988. 140. Engleman, H.M., Kingshott, R.N., Wraith, P.K., Mackay, T.W., Deary, I.J. and Douglas, N.J., Randomized placebo-controlled crossover trial of continuous positive airway pressure for mild sleep apnea/hypopnea syndrome, Am. J. Respir. Crit. Care Med. 159, 461–467, 1999. 141. Hack, M., Davies, R.J., Mullins, R., Choi, S.J., Ramdassingh-Dow, S., Jenkinson, C. and Stradling, J.R., Randomised prospective parallel trial of therapeutic versus subtherapeutic nasal continuous positive airway pressure on simulated steering performance in patients with obstructive sleep apnoea, Thorax 55, 224–231, 2000. 142. Jenkinson, C., Davies, R.J., Mullins, R. and Stradling, J.R., Comparison of therapeutic and subtherapeutic nasal continuous positive airway pressure for obstructive sleep apnoea: A randomised prospective parallel trial, Lancet 353, 2100–2105, 1999. 143. Lamphere, J., Roehrs, T., Wittig, R., Zorick, F., Conway, W.A. and Roth, T., Recovery of alertness after CPAP in apnea, Chest 96, 1364–1367, 1989. 144. He, J., Kryger, M.H., Zorick, F.J., Conway, W. and Roth, T., Mortality and apnea index in obstructive sleep apnea. Experience in 385 male patients, Chest 94, 9–14, 1988.
422
Almoosa and Javaheri
145. Keenan, S.P., Burt, H., Ryan, C.F. and Fleetham, J.A., Long-term survival of patients with obstructive sleep apnea treated by uvulopalatopharyngoplasty or nasal CPAP, Chest 105, 155–159, 1994. 146. Rapoport, D.M., Garay, S.M., Epstein, H. and Goldring, R.M., Hypercapnia in the obstructive sleep apnea syndrome. A reevaluation of the ‘Pickwickian syndrome’, Chest 89, 627–635, 1986. 147. Masa, J.F., Celli, B.R., Riesco, J.A., Hernandez, M., Sanchez, D.C. and Disdier, C., The obesity hypoventilation syndrome can be treated with noninvasive mechanical ventilation, Chest 119, 1102–1107, 2001. 148. Sanders, M.H. and Kern, N., Obstructive sleep apnea treated by independently adjusted inspiratory and expiratory positive airway pressures via nasal mask. Physiologic and clinical implications, Chest 98, 317–324, 1990. 149. Waldhorn, R.E., Nocturnal nasal intermittent positive pressure ventilation with bi-level positive airway pressure (BiPAP) in respiratory failure, Chest 101, 516–521, 1992. 150. Piper, A.J. and Sullivan, C.E., Effects of short-term NIPPV in the treatment of patients with severe obstructive sleep apnea and hypercapnia, Chest 105, 434–440, 1994. 151. de Miguel, J., Cabello, J., Sanchez-Alarcos, J.M., Alvarez-Sala, R., Espinos, D. and Alvarez-Sala, J.L., Long-term effects of treatment with nasal continuous positive airway pressure on lung function in patients with overlap syndrome, Sleep Breath 6, 3–10, 2002.
Part III Clinical Pharmacology
13 Pain Management and Regional Anesthesia
PETER L. BAILEY and RAJBALA THAKUR University of Rochester School of Medicine and Dentistry Rochester, New York
I.
Introduction
Pain may be acute, chronic, or malignant. In this chapter, we focus on how acute postoperative pain and its management affects respiratory function and pulmonary outcome. The goal of optimal pain treatment is justifiable purely on humanitarian principles; however, controlling pain well can be difficult. Over the years, attaining optimal or even good analgesia in patients has remained an elusive goal [1–4]. Numerous elements contribute to suboptimal pain management—including attitudes of health care providers concerning the risks and benefits of pain treatment, assessment methods [5], and technical challenges with epidural analgesia [6–8]. Clinicians remain very concerned about prescribing opioid analgesics because of perceived risks of opioid-induced respiratory depression and addiction. These failures have clear significance: patients have a right to effective pain control and it is likely that optimal pain relief contributes to reductions in morbidity after major surgery [9]. In addition, although the long-term consequences of poorly controlled pain after surgery are not well understood, they are probably significant. Few studies evaluate outcomes 425
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such as intermediate- and long-term functional status as they relate to pain and its management after surgery. The incidence and impact of chronic pain syndromes that develop and persist after surgery can be significant [10], and preliminary evidence suggests that optimal perioperative pain control can reduce such problems. It is well known that pain and impaired respiratory mechanics after upper abdominal or thoracic surgery can result in poor lung function (e.g., impaired gas exchange, atelectasis, hypoxemia) and respiratory complications (e.g., pneumonia) [11]. However, pain is only one of the factors contributing to impaired pulmonary function after abdominal or thoracic surgery. Surgical trauma can elicit viscerally mediated reflexes, which may inhibit normal respiratory function [12]. In addition, numerous factors hinder the ability to eliminate pain in many patients, including the fear that aggressive pain relief strategies carry risks that may outweigh the desired benefits. Analgesic drugs, especially opioids, can negatively and profoundly affect breathing. Understanding the risks and benefits of analgesic therapies and how to administer the drugs judiciously allow clinicians to better achieve optimal pain relief. Our goals for this chapter are to review: (1) the effects of pain, surgery, and surgical trauma on respiratory function; (2) the independent effects of commonly used analgesic drugs and techniques on respiratory function, and (3) the evidence concerning the impact of pain management on respiratory-related morbidities and patient outcomes after surgery. II.
Postoperative Respiratory Dysfunction
The respiratory effects of anesthesia are profound (Figure 13.1). With the use of modern short-acting anesthetic agents and approaches, such effects frequently do not persist more than 1–2 h postoperatively [13]. However, both patient pathology (e.g., lung disease, obesity, obstructive sleep apnea) and surgery can keep pulmonary function from returning to baseline shortly after emergence from anesthesia. Factors underlying or contributing to early postoperative respiratory dysfunction include hypoventilation, ventilation/perfusion mismatch, right to left shunt, depressed cardiac output, and increased oxygen consumption due to shivering [14]. In addition, persistent residual anesthetic drug action can significantly impair respiratory function. Thus, residual neuromuscular blockade can contribute to acute airway obstruction and subsequent respiratory complications, most often in the immediate recovery phase after anesthesia [15]. Phasic activity of the airway muscles of respiration also can be impaired by other anesthetic drugs. Persistent opioid action can compromise ventilatory drive and breathing immediately after surgery.
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Figure 13.1 Schematic illustration of mechanism of physiopathologic effects of postoperative pain (Data from Ref. 349).
Aside from acute and immediate respiratory problems (most often corrected in the post-anesthesia care unit), relatively healthy patients undergoing extremity or superficial surgery do not usually have persistent respiratory problems which can be exacerbated by pain or ameliorated by usual pain control strategies. Thus, after superficial or extremity surgery, most general anesthesia-related pulmonary dysfunction returns to baseline early in the postoperative period and is not likely to play a significant role in postoperative pulmonary morbidity [16]. For example, after lower abdominal surgery there is minimal to no effect on breathing patterns [17] with any analgesic regimen including parenteral/epidural opioids or epidural local anesthetics. Regional anesthesia per se has been associated with improved patient outcomes when compared with general anesthesia [18]. Rodgers et al. [19] reported an overview of randomized trials (142 trials, 9559 subjects) comparing neuraxial (epidural or spinal) versus general anesthesia. The authors found neuraxial anesthesia reduced overall mortality by 33%, deep venous thrombosis by 44%, pulmonary embolism by 55%, transfusion by 50%, pneumonia by 39%, and respiratory depression by 59%. Myocardial infarction and renal failure were also reduced. It remains
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Bailey and Thakur Table 13.1 Alterations in Pulmonary Function and Postoperative Pulmonary Complications after Surgery Alterations in Pulmonary Function Decreased lung volumes Vital capacity (VC) Functional residual capacity (FRC) Atelectasis Decreased expiratory flow volumes and effort Depressed cough Impaired handling of secretions Retention of mucus plugs Decreased compliance Disturbance of sleep breathing patterns Postoperative Pulmonary Complications Ventilation and perfusion abnormalities Shunting Hypoxemia Bronchospasm Pneumothorax Hemothorax Pleural effusions Pneumonia Respiratory failure
unclear if such benefits can also be achieved in relatively healthy patients undergoing peripheral or superficial surgery [20,21]. The respiratory effects of regional anesthetic or analgesic techniques, as well as those of analgesic drugs, both of which can have their action prolonged into the postoperative period, will be considered in separate sections below. The following section focuses on the surgical causes of respiratory dysfunction that persist into the postoperative period. A significant portion of surgical patient morbidity and mortality, especially after major operations, is due to postoperative pulmonary dysfunction. The incidence of such problems can be as high as 10–20% and can be greater than the incidence of cardiac complications, although the latter has received more attention [22]. Postoperatively, patients develop a spectrum of respiratory problems (Table 13.1) ranging from mild hypoxemia and atelectasis, to bronchospasm and pulmonary aspiration, to respiratory failure. Understanding the mechanisms of pulmonary dysfunction after surgery (Figure 13.2) will better allow the prescription and assessment of analgesic therapies tailored to reduce such problems as well as to provide analgesia and comfort.
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Incision effect Expiratory intercostal and abdominal muscle tone
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Figure 13.2 Proposed mechanisms of postoperative pulmonary complications (Data from Ref. 25).
Surgical trauma, especially upper abdominal surgery or thoracotomy, impairs respiratory function through several mechanisms. (1) Direct trauma to, or functional disruption of, respiratory muscles occurs. Muscles that are cut or retracted during thoracotomy include the latissimus dorsi, serratus anterior, pectoralis major and intercostal muscles [23]. (2) Injury also results from spreading, and sometimes fracturing, of ribs, excision of rib periosteum, avulsion of costotransverse ligaments, and intercostal nerve damage. Chest tubes or drains also contribute to trauma and pain. (3) Surgical trauma also causes reflex inhibition via vagal and other inputs, resulting in decreased phrenic nerve output [12] (Figure 13.3). Thus, both voluntary and involuntary inhibition of effective breathing result. Of note, good pain relief may not normalize lung function, due in part to the respiratory depressant properties of opioids (see below) administered for analgesia [24]. In addition, visceral reflexes, which can inhibit diaphragmatic function, can persist despite good pain control [25]. In both animal models and human subjects, shortening of the diaphragm is impaired for days after thoracic [26] or abdominal surgery [27]. Postoperative diaphragmatic dysfunction is intrinsic (e.g., impaired shortening) as well as extrinsic (e.g., decreased phrenic motoneuron
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Figure 13.3 Factors producing respiratory muscle dysfunction after surgical trauma. From left to right: (1) surgical trauma stimulates central nervous system (CNS) reflexes mediated by both visceral and somatic nerves that produce reflex inhibition of the phrenic and other nerves innervating respiratory muscles; (2) mechanical disruption of respiratory muscles impairs efficiency, and (3) pain produces voluntary limitation of respiratory motion. These factors all tend to reduce lung volumes and can produce hypoventilation and atelectasis (Data from Ref. 12).
output). This dysfunction apparently persists even with opioid analgesia [25]. Such postoperative dysfunction is present during quiet tidal breathing as well as during maximal inspiratory efforts. There are marked increases in abdominal and lower intercostal muscle activity after upper abdominal surgery, leading to increased intraabdominal
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pressure and decreased compliance of the abdominal cavity. Activity of the abdominal muscles can reduce lung volumes, both directly because of their insertion on the lower rib cage and indirectly by displacing the relaxed diaphragm cranially secondary to increases in intraabdominal pressures. Epidural analgesia can reverse these changes (see below). The functional residual capacity (FRC), a descriptor of resting lung volume at the end of a normal exhalation, is decreased postoperatively. This reduction is one of the most important pulmonary abnormalities after surgery [13,28]. Other surrogate measures of lung function such as the forced expiratory volume in 1 sec (FEV1), the forced vital capacity (FVC), and the peak expiratory flow rate (PEFR) are also decreased, but these may not be useful predictors of postoperative pulmonary complications [11]. Decreases in FRC augment airway closure, increase atelectasis and shunt, and are a major cause of postoperative hypoxemia. Administration of 100% oxygen, because of absorption atelectasis, also has been demonstrated to worsen atelectasis [29,30]. Decreased FRC is widely noted with almost all general anesthetic techniques, regardless of the type of surgery. Certainly such lung volume changes are a greater issue in patients with concomitant problems such as obesity, underlying lung disease, and advanced age. Surgical site also impacts FRC and vital capacity, and the time course of this impact is significant (Figures 13.4 and 13.5). After upper abdominal surgery, FRC decreases by about 70% at 24 h and gradually returns to normal by day 7–10 [28]. A common clinical sign of impaired respiratory function and/or failure is rapid shallow breathing. Tachypnea itself is one of the better predictors of impending respiratory failure. After abdominal or thoracic surgery, a restrictive pattern with reductions in inspiratory capacity, vital capacity, FVC, PEFR, maximum minute ventilation and FRC is typical [31]. Such dysfunction can last for days to more than a week and is commonly referred to as splinting. Patients are unable and/or unwilling to breathe deeply. Surgical trauma and pain can inhibit coughing that leads to retention of secretions with lobular or lobar collapse [32]. Such changes in respiratory function are much less common after lower abdominal surgery [27] and usually minimal after superficial or extremity surgery. The adverse effects of surgery are most pronounced after upper abdominal or thoracic surgery [22]. Major abdominal surgery is followed by moderate hypoxemia for up to two weeks and by episodes of severe episodic desaturations both in the immediate and late postoperative periods [25,33]. As discussed above, disturbances in pulmonary mechanics and chest wall function are due to many factors. Vital capacity is reduced to less than half of preoperative values after surgery requiring sternotomy and requires a week to return to near baseline levels [34]. Open procedures with novel approaches, minimally invasive, which seek to reduce
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trauma to structures involved in respiratory function, can reduce postoperative pulmonary dysfunction [35]. Pulmonary morbidity can range from 2% after minor surgery to as high as 33% after major operations [36]. Minimally invasive procedures (e.g., laparoscopy, thoracoscopy) do not impair pulmonary function to the degree that open procedures do [37,38].
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Figure 13.5 Vital capacity (VC) (mean SEM) expressed as a percentage of preoperative value for the two groups. (g) ¼ Lower abdominal surgery; (f) ¼ upper abdominal surgery. Significant difference from control: *P 5 0.05; **P 5 0.01. Significant difference between the two groups: yP 5 0.01 (Data from Ref. 27).
III.
Other Effects of Pain and Surgical Trauma
Pain can influence respiratory function indirectly through a wide range of effects. These will be reviewed briefly in the following sections to highlight how respiratory function, through its links to other physiological systems, can be additionally stressed after surgery and by painful conditions. A. Effects of Pain on the Stress Response
Nociception and surgical trauma can evoke a complex sequence of neuroendocrine and inflammatory responses [39]. This stress response is mediated via the somatosensory and sympathetic pathways and is proportional to the extent of injury [40]. Nociceptive impulses arise from the site of injury, enter the neuraxis predominantly in the dorsal horn of the spinal cord, and if not blocked, continue on to the brain via the spinothalamic and spinoreticular pathways (Figure 13.6). Over time, local and descending activity modulates the pain response. Elevated sympathetic
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Figure 13.6 Neural pathways for sympathetic activation in response to surgery. Painful stimuli transmitted through afferent nociceptive pathways activate sympathetic efferent pathways. Sympathetic stimulation of the myocardium increases heart rate (HR) and inotropy. Stimulation of the peripheral vascular bed produces vasoconstriction with a resulting increase in blood pressure (BP). EPI ¼ epinephrine; NEPI ¼ norepinephrine (Data from Ref. 25).
efferent activity and the overall stress response become manifest by increases in serum levels of epinephrine, norepinephrine, growth hormone, cortisol, renin, aldosterone, C-reactive protein and antidiuretic hormone. A broad spectrum of postoperative complications is postulated to be exacerbated by inadequately controlled acute pain (Table 13.2). Tendencies toward a hyperdynamic circulation, increased metabolism and oxygen consumption, and decreased respiratory function in postoperative patients combine to produce decreased physiologic reserve margins and push compensatory mechanisms to their limit [41]. Adverse metabolic and endocrine effects may also be exacerbated if not caused by severe pain. This includes increased glycolysis, lipolysis, and proteolysis. The catabolic response to surgery can be enhanced, as can sodium and water retention due to increased cortisol, epinephrine, antidiuretic hormone and aldosterone secretion [42]. Negative nitrogen balance problems will be exacerbated by uncontrolled pain, which causes immobility, poor appetite and nutrition.
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Table 13.2 Potential Adverse Effects of Postoperative Pains General Immobility Increased length of stay Respiratory Splinting Atelectasis Decreased lung volumes and function Hypoxia Pneumonia Respiratory failure Cardiovascular Hypertension Tachycardia Arrhythmias Myocardial infarction, failure Hemostatic Hypercoagulable state Venous thrombosis Coronary artery thrombosis Metabolic/Endocrine Increased oxygen consumption Poor nutrition Negative nitrogen balance Fluid and electrolyte imbalance Hyperglycemia Insulin resistance Lipolysis Immunological Immunoincompetence Psychological Depression Anxiety Failure to thrive
The postoperative stress response is now thought to be largely maladaptive and to play a major role in complications, morbidity and mortality [43]. Analgesic management, which is pre-emptive, multimodal, and includes neuraxial blockade with local anesthetics, will reduce predominantly endocrine and metabolic responses. Neuraxial analgesia can also result in less immunomodulation and fewer infections when compared with systemic opioid analgesia [44,45]. However, it is through the use of less traumatic surgical techniques (e.g., minimally invasive surgery) that the inflammatory responses will be reduced [21].
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A few studies suggest that reduction of the stress response can improve patient outcome [20,46]. Brodner et al. [47], in a randomized trial, included assessment of the stress response in an outcomes study of three groups of patients undergoing radical cystectomy. Group one received general anesthesia, group two received general anesthesia combined with epidural anesthesia, and group three was the same as group two, but with postoperative patient-controlled epidural analgesia, early oral nutrition and enforced ambulation. Group three patients had improved dynamic pain relief and better recovery of bowel function as well as signs of reduced stress response and improved metabolism. Only 15 patients were in each group and no other significant outcome differences were found. More studies proving outcome improvement due to stress response reduction are needed. It remains unclear if the stress response can serve as a meaningful surrogate marker for outcomes. In addition, it is not well defined which aspects of the stress response are most important to control in order to improve outcome. The direct link between adverse outcomes and pain per se also remains largely hypothetical. However, therapies designed to optimally control pain are intimately linked to the reduction of the consequences of surgical trauma, such as the stress response. Studies have determined that certain postoperative analgesic techniques, most notably epidural analgesia, can produce benefits (see below) [11,43]. Unfortunately, acceptance of the importance of pain control in reducing the postoperative stress response and improving patient outcome has been limited. Rigg et al. have noted that anesthesiologists and surgeons are still divided about the value of epidural analgesia [6]. Current surgical treatises usually make little reference to the role of pain control. It is only with persistence, additional studies and allocation of adequate resources that the benefits and need for optimal pain control will gain recognition. B. Effects of Pain on Pulmonary Function
Acute pain, especially after upper abdominal or thoracic surgery, can impair respiratory mechanics and breathing. Splinting of the chest and abdominal walls decreases tidal volume, vital capacity, FRC and ventilation [48,49]. Deep breathing and coughing are suppressed. A restrictive pattern develops with reduced lung volumes and increased respiratory rate; FRC frequently is below closing capacity. These changes predispose patients to alveolar collapse, hypoxemia and impaired gas exchange. Impaired coughing and retention of secretions also contribute to atelectasis, fever, pulmonary infection, and sepsis. Any or all of these factors can culminate in respiratory failure. As mentioned, pain relief is not necessarily sufficient to normalize pulmonary function after surgery. Pain relief, achieved solely with opioids
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after abdominal surgery, while suggested by some to reverse postoperative pulmonary dysfunction (as measured by FEV1), has not been found to do so by others [50], perhaps due to the different measures of pulmonary function used [51]. Manikian et al. [51] have demonstrated that pulmonary dysfunction after upper abdominal surgery can be ameliorated by thoracic epidural analgesia with local anesthetic. This is consistent with the hypothesis that inhibitory reflexes, initiated by visceral irritation and acting on the phrenic nerve–diaphragm unit, may play a key role in postoperative respiratory dysfunction. C. Effects of Pain on Cardiovascular Function
Cardiovascular morbidity is the leading cause of death after surgery [52]. Interestingly, most episodes of myocardial ischemia are silent and do not produce significant hemodynamic changes [31,53]. Pain can stress directly and indirectly the cardiovascular system, leading to coronary artery vasoconstriction [54]. Stimulation of the sympathetic nervous system causes tachycardia, hypertension and increased systemic vascular resistance. All of these factors increase cardiac work and myocardial oxygen demand and can lead to or exacerbate myocardial ischemia. Myocardial infarction or failure can result [55], in turn further augmenting sympathetic activation resulting in a vicious cycle. Concomitant respiratory effects mentioned above can further worsen cardiac conditions by impairing oxygen uptake and delivery. Hypercapnia also can further stimulate the sympathetic nervous system and cause pulmonary hypertension, right ventricular failure and even right-to-left shunting if a patent foramen ovale exists. Effective analgesia may reduce postoperative myocardial ischemia and cardiovascular instability [52,56,57]. Recently, Beattie et al. [58] reported the results of a meta-analysis determining whether postoperative epidural analgesia continued for more than 24 h reduced postoperative myocardial infarction or death. The authors found 17 studies, of which 11 were randomized controlled trials comprising 1173 patients. Postoperative epidural analgesia was confirmed to provide superior pain control and the rate of postoperative MI differed by a significant amount (3.8%). Subgroup analysis revealed thoracic epidural analgesia produced an even greater, statistically significant reduction (5.3%). Thromboembolic phenomena are a serious postoperative problem. Sympathetic activation in the postoperative period is also associated with a hypercoagulable state, which can increase the risk of coronary artery thrombosis and associated coronary events [59]. Surges in catecholamines and angiotension can decrease regional and lower extremity perfusion [60]. Decreased ambulation also increases thromboembolic tendencies. Effective analgesia, with local anesthetic resulting in a degree
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of sympathetic blockade, improved extremity blood flow and reduced hypercoagulability [61–63]. D. Other Effects of Pain
Adverse gastrointestinal effects induced by acute pain include decreased peristaltic activity and motility in the gut as well as increased intestinal smooth muscle and sphincter tone. Ileus and intestinal dilatation are frequently experienced in the postoperative period [64]. Ileus is exacerbated by opioid analgesics; ambulation helps regain normal bowel function [25]. Neuraxial analgesia fosters more rapid return of bowel function when compared with systemic opioid analgesia, and dynamic pain control is essential for early ambulation after surgery [65,66]. Decreased renal and splanchnic blood flow leads to decreased glomerular filtration, resulting in retention of water and sodium. Urinary retention may also be exacerbated by pain-induced autonomic nervous system effects. Sleep deprivation, mental stress, and depression can result from poorly controlled pain [67]. Finally, poorly controlled pain can increase both length of patient hospital stays and health care costs [68]. IV.
Pain Control and Chronic Pain
Central and peripheral sensitization, mechanisms integral to the development of chronic pain, begin to develop soon after the onset of acute pain. Tissue trauma and the subsequent production or release of numerous substances such as prostaglandins, histamine, and bradykinins contribute to both acute and subacute pain states. Pain starts a cascade of responses that can evolve into intractable chronic pain states. Neurotransmission related to pain, if prolonged, leads to expanded receptive fields, increased sensitivity of pain-responsive neural elements and permanent changes in pain-modulating neuronal systems in the spinal cord, leading to persistent postoperative neuropathic pain [69]. How frequently postoperative pain develops into a chronic pain condition and whether certain analgesic approaches can ameliorate such problems are inadequately studied. Up to 50% of thoracotomy patients have persistent pain six months or more after surgery [70]. Effective perioperative analgesia can significantly reduce the incidence of persistent pain after thoracotomy [71]. V.
Effects of Pain and Pain Management on Respiratory Function
It is widely held that pain stimulates respiration. It is also believed that pain or strong stimulation can counter troublesome opioid-induced
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respiratory depression. Pain pathways originating in the periphery have numerous connections and projections within the CNS [72]. Pain can stimulate breathing through general arousal mechanisms and through subcortical mechanisms [73], and it seems to re-set resting ventilation by increasing tidal volume and respiratory rate, without affecting chemoreflexes [74,75]. Borgbjerg et al. [76], in an experimental model of pain in human volunteers, found no effect of pain on the slope of the ventilatory response to carbon dioxide. Sarton et al. [75] confirmed that experimental pain in human volunteers stimulated ventilation through an increase in tonic ventilatory drive mechanisms that were independent of chemoreflexes. On the other hand, others have found that the relief of pain in humans who had suffered upper extremity trauma decreased both minute ventilation and the slope of the ventilatory response to CO2 [77]. Pain management can entail a wide array of techniques and drugs. An almost infinite number of strategies can be employed, if one considers not only the drugs utilized, but also the timing, route and method of administration. Add to this the many drug combinations that can be administered in an attempt to achieve optimal ‘balanced’ analgesia, and it becomes apparent that a complete review is not possible. In this chapter we focus on the respiratory effects of: (1) opioid agonists as the standard parenteral analgesic; (2) regional analgesia with local anesthetic as the standard agent used for pain relief with nerve blockade, and (3) other miscellaneous agents which continue to be explored as analgesics. These miscellaneous analgesic agents will be considered separately, regardless of their route of administration, to provide an overview of their actions, applications, and how their administration affects breathing. Opioids remain the most potent and commonly prescribed analgesic. Despite significant advances in opioid pharmacology, the holy grail of pain control—excellent analgesia without respiratory depression—remains elusive. Hope in achieving this goal, brought on by the suggestion that different mu receptors account for analgesia and respiratory depression, has not been fulfilled [78]. We are still unable to separate the gold standard intense analgesia that opioids can produce from either commonly associated side effects (e.g., pruritis) or the uncommon but most feared adverse effect (i.e., respiratory depression) intrinsic to analgesic approaches which use these drugs. Accurate assessment of pain also remains problematic. The measurement of pain relief is often done with a visual analogue scale as a means of quantifying pain. However, non-uniform approaches to quantifying pain contribute to the variability found in pain studies, pain assessment, and pain relief. Importantly, although now recognized as the true measure and goal of optimal pain relief, dynamic pain scores (pain rating during motion and motor function) also are not consistently determined or widely used. It is only with dynamic pain relief that a patient
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can perform the activities, such as deep breathing, coughing, and ambulation, which permit some of the benefits of analgesia to be achieved. A. Opioids
The opioid receptor was identified in nervous tissue in 1973 [79–81]. Endogenous opioid peptides act at three receptors (mu, kappa, and delta) that are involved in the physiologic responses to pain, emotion, and stress. Endogenous opioid systems also have other physiological roles, including cardiovascular, thermoregulatory, and immune effects. Importantly, endogenous opioids are found in high concentrations in areas of the peripheral and central nervous system that play a role in respiratory drive and function. Endogenous opioid peptides are widely distributed in brainstem nuclei regulating respiration [82]. High concentrations of opiate receptors have been found in many supraspinal brain respiratory centers including the nucleus tractus solitarius, nucleus retroambigualis and nucleus ambiguous [83]. Specific chemosensitive brain areas also mediate opioid-induced respiratory effects [84]. The mu opiate receptor mediates most clinically important analgesia. Investigations utilizing opiate receptor knockout mice demonstrate that morphine-induced analgesia is mu receptor-mediated [85]. The role(s) of the delta receptor in analgesia and respiratory control, if any, remains unclear. Mu-1 receptors also appear more prominent in supraspinal analgesia, whereas delta-receptors may be more important for spinal analgesia [86]. Recent studies comparing the effects of opioids in mice with the mu opioid receptor (wild-type) versus mice without the mu opioid receptor (knockout mice) confirm and define the role of endogenous and exogenous opioids in the control of breathing. Resting ventilation was slightly increased (15%) in the knockout mice, but the ventilatory response to CO2 did not differ between the two groups, suggesting only a minor role for the mu opioid receptor in the control of breathing. Morphine doses in normal mice, which caused analgesia as well as depression of resting ventilation and the slope of the ventilatory response to CO2, produced no analgesia or respiratory depression in knockout mice (Figures 13.7 and 13.8). Naloxone increased resting ventilation and the slope of the CO2 response in the wild-type and the knockout mice, suggesting a role for endogenous opioids in the control of breathing. The mu opioid receptor was confirmed to mediate both analgesia and respiratory depression induced by exogenous opioid administration [87]. Of note, mu receptors are located in both the brain and spinal cord with highest concentrations in the periaqueductal gray and substantia gelatinosa, respectively. Mu receptor opioid-induced analgesia is
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dose-dependent. Pharmacological distinction of an analgesic mu receptor (mu1) and a respiratory depression mu receptor (mu2) is suggested but remains in question [88]. Kappa-receptor activation may explain in part some of the effects (analgesia with limited respiratory depression) of some mixed agonist–antagonists such as nalbuphine. At least three separate kappa receptor subtypes have been isolated. Kappa-3 receptors are of particular interest because of their high density within the brain and association with supraspinal analgesia, whereas other kappa receptors relate to spinal analgesia [89]. Specific opiate receptor mRNA expression can be found in the CNS and correlates well with opiate binding in major ascending and descending pain pathways [90]. These include the amygdala, the mesencephalic reticular formation, the periaqueductal gray matter (PAG), and the rostral ventral medulla. Opioid actions in the PAG influence, through direct neural connections, the rostral ventromedial region of the medulla (RVM). The substantia gelatinosa of the spinal cord possesses a dense collection of opiate receptors [91]. The documentation of opiate receptors in the spinal cord was a landmark discovery; direct application of opioids to these regions creates intense analgesia. Opioid receptors are also found in close association with peripheral chemoreceptors in the carotid body [92]. Respiratory Actions of Opioids
The respiratory depressant actions of opioids represent their single most serious adverse effect [93]. There is no more dreaded, or conceivably avoidable, complication than that of a respiratory arrest due to a pain control method with opioids gone awry. Although significant adverse events related to opioid-induced respiratory depression are presumably preventable, numerous studies document that they persist with a perioperative incidence of approximately 1%, no matter what the route of drug administration. Opioids do produce some respiratory actions that may be desirable or therapeutic. Pain and/or anxiety can induce excessive spontaneous ventilation, resulting in respiratory alkalosis and excessive but potentially ineffective work of breathing. Opioids, by decreasing both pain and central ventilatory drive, are effective agents in such conditions. The lack of adequate pain relief can also cause postoperative respiratory dysfunction. Appropriate opioid analgesia can improve synchronous breathing and decrease voluntary muscle tone, resulting in improved dynamic total respiratory compliance in awake but mechanically ventilated intensive care patients [94]. The antitussive actions of opioids are well known. Opioids are also excellent agents for depressing upper airway, tracheal, and lower respiratory tract reflexes. The drugs can blunt the pulmonary
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vasoconstrictive response to tracheo-bronchial stimulation that can occur with suctioning [95]. The exact mechanisms underlying these clinical observations are not clear. Opioid receptors have been demonstrated in smooth muscles of the trachea and bronchi, as well as alveolar walls, but not in the small airways [96]. Opioids blunt or eliminate somatic and autonomic responses to tracheal intubation. The drugs allow patients to tolerate endotracheal tubes without coughing or bucking, which when pronounced can lead to decreased lung volumes, atelectasis, hypoxemia, hypercarbia, hemodynamic instability, and other deleterious effects. Impaired gas exchange results from the marked disturbance in ventilatory pattern as well as from loss of lung volume that bucking produces. Opioids have been used in the treatment of acute asthma for decades, relieving dyspnea and other symptoms of respiratory distress associated with conditions such as asthma and heart failure. Fentanyl also has antimuscarinic, antihistaminergic, and antiserotonergic actions and may be more effective than morphine in patients with asthma or other bronchospastic diseases. Certain intravenous agents, including opioids, minimally alter pulmonary gas exchange [97]. The minimal impact of opioids on hypoxic pulmonary vasoconstriction (in contrast to the potent inhalation agents and other potent vasodilators such as nitroprusside), coupled with associated hemodynamic and bronchomotor stability, contribute to the minimal interference with pulmonary gas exchange observed after opioids and many other intravenous anesthetics. All mu-receptor stimulating opioids cause dose-dependent depression of ventilation in humans, primarily through a direct action on brain stem respiratory centers. Controversy persists whether different subclasses of mu opioid receptors have disparate roles in opioid-induced respiratory depression [98]. Opioids interfere with pontine and medullary respiratory centers that regulate respiratory rhythmicity. In the cat, different medullary inspiratory neurons have varying sensitivity to fentanyl. Both dorsal (DRG) and ventral (VRG) respiratory groups are affected. In the DRG, a continuous discharge has been reported to replace rhythmic discharge after fentanyl; this change was reflected in phrenic nerve activity. In the VRG, inspiratory neuron activity was completely abolished. These effects can lead to sustained contraction of respiratory muscles and cessation of respiration [99] (Figure 13.9). The stimulatory effect of CO2 on ventilatory drive is also significantly reduced by opioids, thus decreasing the slopes of the ventilatory and occlusion pressure responses to CO2, and shifting to the right minute ventilatory responses to increases in PaCO2. In addition, the apneic threshold and resting end-tidal PCO2 are increased by opioids.
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Control
9 min after Fentanyl 50 mg/kg IV
15 min after Fentanyl
45 min after Fentanyl
5 sec Figure 13.9 Effect of fentanyl (50 mg/kg) on an inspiratory unit in the ventral respiratory group. The unit’s activity is totally abolished by fentanyl. Full recovery occurred in 45 min. The effect of fentanyl on this unit is different from that on the units in the dorsal respiratory group (Data from Ref. 99).
Intravenous or intrathecal opioids also decrease hypoxic ventilatory drive, due to peripheral effects at the carotid body and to direct central effects [100,101]. Hypoxic drive can be blunted or eliminated by low, analgesic doses of opioids. Hypoxic ventilatory decline, the delayed decrement in minute ventilation that occurs after the peak ventilatory response to acute hypoxia when hypoxia persists, may be potentiated by opioids [102]. Opioids also blunt the increase in respiratory drive normally associated with increased loads such as increased airway resistance [103]. Differences in opioid-induced respiratory depression between males and females have been reported [104]. Analgesic doses of IV morphine have been reported to depress ventilation in both sexes but by different
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mechanisms: ventilatory CO2 and hypoxic sensitivities were depressed only in women whereas CO2 and hypoxic responses were shifted only in males. Opioid-induced effects on the control of respiratory rhythm and pattern include increased respiratory pauses, delays in expiration, irregular and/or periodic breathing, reductions in sighs, and decreased, normal, or increased tidal volume. The prolonged expiratory time in the respiratory cycle induced by opioids frequently results in greater reductions in respiratory rate than tidal volume. Fentanyl depresses respiratory drive, phase timing, and activation of respiratory muscles [105]. Sighs and normal variation in tidal volumes can be significantly reduced, contributing further to opioid-induced respiratory problems. With systemically administered agents, the clinical duration of analgesia coincides with the risk of respiratory depression. Although recurrence of respiratory depression can occur with systemically administered opioids such as fentanyl, it is uncommon [106]. Many factors can affect both the magnitude and duration of respiratory depression after opioids (Table 13.3). The concomitant administration of other drugs, such as a benzodiazepine, can markedly potentiate respiratory depression [107]. Looi-Lyons [108] reported a patient who experienced troublesome respiratory depression 6 h after a general anesthetic that included 15 mg of midazolam. Midazolam can have a prolonged effect after such doses, especially if hepatic metabolism is impaired. Drugs which either directly depress ventilation, or which produce sedation, can have strong synergistic respiratory depressant actions when combined with opioids. Patients who are sleeping are usually more sensitive to the depressant effects of opioids. Even small doses of opioids markedly potentiate the normal right shift of the PaCO2–alveolar ventilation curve that occurs Table 13.3 Factors Affecting Opioid-Induced Respiratory Depression after Surgery 1. 2. 3.
4. 5. 6. 7. 8. 9. 10. 11.
Dose Method of administration Increased brain penetration/drug delivery a. Decreased distribution (decreased cardiac output) b. Increased unionized fraction (respiratory alkalosis) Decreased re-uptake from the brain (respiratory alkalosis) Decreased clearance (decreased hepatic blood flow) Active metabolite accumulation (morphine, meperidine; renal failure) Secondary peaks in plasma opioid levels Increased ionized opioid at receptor site (postoperative respiratory acidosis) Sleep Extremes of age Metabolic alkalosis
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during natural non-REM sleep. For several days postoperatively, sleep is associated with hypoxemia [109]. The mechanisms and implications of postoperative sleep disturbances are complex, and relate not only to respiratory problems, but also to cognitive and hemodynamic disturbances [110]. Both sleep and morphine relatively spare the diaphragmatic but decrease the thoracic (ribcage) component of breathing [111]. Sleep also impairs tonic and phasic upper airway muscle activity that accompanies breathing [112]. Sleep disorders (e.g., obstructive sleep apnea) can place patients at greater risk for troublesome respiratory depression related to opioids [113]. Sleep disorders of breathing include increased upper airway resistance, obstructive sleep apnea, hypoventilation, central apnea, and Cheyne-Stokes breathing [113]. Acute opioid therapy may not contribute to disordered breathing in adults without sleep apnea [114], but in patients with sleep disorders, opioid therapy can produce apneic and hypoxic episodes which are more severe in non-REM sleep, irregular breathing patterns, pronounced obstructive hypoventilation, and severe hypoxia. CPAP may be ineffective in ameliorating these abnormalities [113]. It is thought that since the carotid bodies are primarily responsible for the recognition and response to airway occlusion and the breath-to-breath control of ventilation, carotid body dysfunction and/or disruption of the processing of information from them underlie opioid-induced sleep-disordered breathing. However, even when pain is well controlled and opioids are avoided, profound postoperative sleep disturbances occur [115]. It is beyond the scope of this chapter to review the impact of conditions and diseases that can affect opioid pharmacology. Clinicians should recognize that in some conditions, certain opioids should be avoided. For example, in renal failure, metabolites of either morphine or meperidine can accumulate and produce excessive respiratory depression or toxicity. Clinicians frequently use decreased respiratory rate as a sign of impending excessive opioid-induced respiratory depression. Opioids can usually be titrated to effect, especially in anesthetized patients, by observing dose-dependent decreases in the spontaneous respiratory rate. However, numerous reports note that respiratory rate, especially in the postoperative setting, cannot serve as a reliable index of the magnitude of opioid-induced respiratory depression. Patients experiencing excessive opioid effects may be apneic or severely hypopneic, but may also remain responsive and often breathe when commanded or stimulated to do so. Respiratory rate is usually slowed drastically if not absent in overt opioid overdose. It is well known that opioids have undesirable gastrointestinal effects, including nausea, vomiting, and decreased motility, effects which can exacerbate postoperative ileus and bowel distention. It has been postulated that such actions lead to greater elevation of the diaphragm after surgery and impairment of pulmonary function [41]. Sedation from opioids, or from
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any other drugs, will also inhibit patients from sitting upright or ambulating. The sedentary supine position in the sedated patient also favors decreased lung volumes and adverse physiological respiratory sequelae such as atelectasis and hypoxemia. Peak onset of respiratory depression after an analgesic dose of morphine is slower than after comparable doses of fentanyl, taking 15–30 min versus approximately 5 min, due in part to the lower lipid solubility of morphine. Due to its low lipid solubility, morphine’s plasma concentrations and onset of action are nearly identical after intravenous and intramuscular administration [116]. Nevertheless, the selection of morphine as an IV analgesic for acute pain control in the immediate postoperative period remains commonplace. Respiratory depression induced by small doses of morphine usually lasts longer than the depression induced by equipotent doses of fentanyl. Intravenous fentanyl (100 and 200 mcg/70 kg) has a faster onset of action and results in a somewhat shorter period of respiratory depression than an equipotent dose of meperidine (65–75 mg/70 kg) [117]. Even though fentanyl has a shorter onset and quicker recovery than morphine and meperidine, small doses (1–2 mcg/kg) can produce very significant respiratory depression, while peak effect of fentanyl occurs 4–6 min after intravenous injection, residual respiratory depression can persist for more than 1 h. Plasma fentanyl concentrations of 1.5–3.0 ng/ml are usually associated with significant decreases in CO2 responsiveness [118]. Intraveously injected sufentanil also has a rapid onset of effect, peaking in 4–6 min. Sufentanil (0.1–0.4 mcg/kg) has been reported to produce shorter lasting respiratory depression and longer lasting analgesia than fentanyl (1.0–4.0 mcg/kg) [119]. Sufentanil is distributed in concentrations of 50 mcg/ml. This, combined with the fact that it is approximately 10 times as potent as fentanyl, has limited its widespread clinical use as an analgesic. Opioid Analgesia
The therapeutic window for all currently used opioid agonists is narrow. For example, the plasma concentration of fentanyl that produces analgesia (1–2 ng/ml) also produces respiratory depression. Hence, as good or optimal analgesia is approached with systemically administered opioids, so are the risks of side and adverse effects. Compounding this matter is the fact that individual variation in fentanyl requirements ranges from 0.2 to 8.0 ng/ml with a log-normal distribution [120]. This variability holds for other opioid agonists as well. Morphine patient-controlled analgesia (PCA) requirements average a few milligrams per hour, but reportedly can range from 0 to 16.5 mg/h [121]. It is not uncommon for patients to require more than 100 mg of morphine within the first 24 h after major surgery.
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Popular opioid agonists, frequently used for acute pain control, include morphine, meperidine, fentanyl, and hydromorphone. These agents have well-defined clinical pharmacokinetcs that provide relatively rapid onset and duration of drug action. Other agonists, such as methadone, can be effective but can produce long-lasting effects (e.g., 20–30 h) [122]. Opioid administration traditionally has been via the intramuscular route. While simple, this approach can cause discomfort, requires multiple nursing interventions, and results in significant peaks and troughs in drug levels. Although certainly still popular in many settings, generally intramuscular opioid injection represents a crude approach to pain control. In addition, patients perceive and rate their pain differently than health care providers such as nurses [123]. This discrepancy contributes to inadequate pain treatment when the administration of analgesia is dependent on an individual other than the patient determining the need for pain medication [2,124]. Pain control will not be optimal or consistently good with intramuscular administration of opioids and because of the potential for greater peaks and troughs in drug levels and analgesia, the risk of respiratory depression may be increased [124]. Patient-controlled analgesia (PCA) with opioids offers patients a degree of control over their pain and usually allows proper titration [125,126]. Safe PCA practice includes the prescription of appropriate unit doses, lockout intervals and background infusions as well as proper patient instruction. Background infusion rates may be effective but are frequently limited, since studies have failed to demonstrate their association with any benefit [127,128]. In addition, the use of such background infusions can increase the risk of respiratory depression, if only because of an increased opportunity for pump programming errors [129]. Lockout intervals for PCA are usually on the order of 5–15 min. This will allow for the full onset of drug action with more rapid-onset drugs such as fentanyl, and the nearly complete onset of action with most other opioids. Safe PCA practice also usually includes limiting the activation of PCA devices to the patient. The best of intentions can have deleterious effects, from excessive opioid action and respiratory depression, when family members or friends activate PCA therapy based on their perceived needs of patients. Instructions on this aspect of PCA care should be clear. Exceptions, such as when parents help with PCA of a child, need to include instructions and guidelines. PCA in pediatric patients has been reviewed and deemed appropriate in children more than five years old [130]. Respiratory depression remains the greatest concern with the use of opioid IV PCA in pediatric patients, with a reported incidence ranging from 0 to 7%. The use of background infusions and/or drug combinations appears to be associated with respiratory depression in children as in adults. Unit doses in adults for meperidine, morphine, hydromorphone and fentanyl are approximately 15, 1.5, .15, and .015 mg, respectively. In many
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settings and patient conditions, a wide range of regimens is required. Obviously, factors such as the type of surgery will also determine the dose and schedule requirements. There is a concentration–effect relationship concerning opioid-induced analgesia and respiratory depression [118]. Fentanyl analgesia is associated with plasma concentrations of 1–2 ng/ml, but significant respiratory depression can occur with fentanyl serum concentrations of 2 ng/ml. When good analgesia is achieved, arterial CO2 partial pressures may be only moderately elevated (48 mm Hg), but apneic episodes may still occur [131]. A plasma concentration of 3–4 ng/ml of fentanyl produces a 50% depression of the slope of the ventilatory response to CO2. Severe respiratory depression is uncommon in association with PCA opioid therapy [132,133] but has been reported [129,134–136]. Overall, the incidence ranges from 0.1% to 1.0%. LooiLyons et al. [108] prospectively studied 4,000 patients who received morphine PCA postoperatively and reported nine cases of troublesome respiratory depression. White [137] categorized problems with PCA into (1) operator errors, (2) mechanical problems, and (3) patient errors. Operator (i.e., human) errors, recently receiving much attention, range from prescription transcription errors (e.g., poor handwriting) to faulty pump programming. Mechanical errors include syringe breakage, defective equipment, and program corruption. Patient errors are largely due to the misunderstanding of PCA usage. All of the above can contribute to troublesome respiratory depression or even death. Clinical detection of significant respiratory depression is limited by the clinical methods commonly available and used. Most often, observation of a patient’s respiratory effort and rate is infrequent (e.g., every 1–2 h) on hospital wards, especially at night. Respiratory rate can be a misleading measure of respiratory depression. Other potentially more effective methods, which might combine telemetry with pulse oximetry, mechanoelectric measurement of chest and abdominal wall motion (e.g., impedance or respiratory inductive plethysmography), or capnography, are not widely used outside intensive care settings nor are they without pitfalls themselves. Frequent (e.g., every 30–60 min) monitoring of respiratory rate and the level of consciousness and responsiveness can be a very good clinical tool, as opioid-induced respiratory depression is usually accompanied by sedation and is not usually sudden in onset. The ideal PCA system would include feedback loops whereby measured respiratory variables would reliably indicate excessive drug effect and simultaneously result in stimulation of the patient to breathe (via some audible and/or mechanical mechanism) and notification of health care providers in a timely manner to the presence of excessive drug effect. Whether or not the low incidence of such problems will merit the development and deployment of such systems in the future remains to be seen.
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Although the potential for adverse respiratory outcomes associated with opioid analgesia has been well highlighted [134,138], little data exists to help determine if the use of PCA opioids in the treatment of postoperative pain can improve outcome. Gust et al. [139] reported that, when compared with a control (nurse-controlled analgesia), both PCA with an opioid alone and opioid PCA plus a nonsteroidal anti-inflammatory drug decreased postoperative atelectasis and produced superior pain control. A small number [120] of patients were studied and no postoperative pneumonias occurred, despite a reported incidence of pneumonia of 3–16% in the patient population similar to that in the study. Wasylak also reported that smaller decreases in postoperative pulmonary function might be achieved with IV PCA, when compared with IM opioid analgesia [140]. Walder et al. published a quantitative systematic review of patient-controlled analgesia [141] in which 32 trials, 22 using morphine, were evaluated. Their findings suggested that in the postoperative setting, PCA with morphine, when compared with conventional analgesia, improved analgesia, was preferred by patients, and potentially decreased pulmonary complications. This last conclusion was based on findings in two of the morphine trials. Other methods of delivering opioids are numerous but not as widely used as PCA. Such therapies still can carry the usual opioid-associated risks, but due to unique factors associated with their design, they can also have additional risks. For example, the effect of transdermal fentanyl can be markedly influenced by blood flow and heat applied to the skin containing the fentanyl patch. Drug depot in the dermis can create prolonged (e.g., 24 h) effect and risk for adverse effects [142]. Other novel methods of delivering opioids include oral, oral transmucosal, nasal, and transpulmonary routes. Kehlet [21] notes that analgesic strategies that rely on opioids alone cannot reliably produce dynamic pain relief. He believes that it should not be anticipated that opioids can or should result in improved outcomes other than pain control. Results of a meta-analysis [11] confirm this. Extremes in Spectrum of Opioid Effect: Tolerance and Overdose
Resistance (e.g., tolerance) to opioid effect develops over time; usually three or more weeks of opioid consumption are required for clinically significant tolerance to develop. The mechanisms underlying the development of tolerance have been reviewed [143]. Acute tolerance to the analgesic effects of opioids such as remifentanil also has been reported. It is unlikely that acute tolerance to opioid-induced respiratory depression develops over the same time frame as tolerance to the analgesic effects of opioids. In rats, acute tolerance to the analgesic actions of opioids occurs rapidly while tolerance to the respiratory depressant actions of opioids may take months to develop [144]. Significant tolerance to the respiratory depressant actions of opioids on hypoxic ventilatory responses can take months to
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develop [83]. Delayed development of tolerance to the respiratory depressant effects of opioids, in the presence of tolerance to the analgesic effects of opioids, likely narrows the therapeutic window of opioids to a precarious degree [145]. Hence, the opioid-tolerant patient with severe pain may continue to experience and complain of pain and require additional opioid doses until opioid-induced apnea occurs. Cross-tolerance to the respiratory depressant actions between different opioids may be incomplete or unpredictable. The more potent opioids, such as sufentanil, because of their greater intrinsic activity and low percent receptor occupancy, result in less tolerance. However, morphine does not appear in general to have a clinically relevant ceiling to its analgesic effects, although the severity of side effects may limit dose escalation. In animals, intrinsic efficacy is inversely related to the propensity for tolerance to develop when opioids are administered by continuous infusion but not by once-daily dosing. Thus, the method and schedule of drug administration may also affect the development of tolerance [146]. Pain relief in the opioid-tolerant individual should be appropriate for the degree of pain he/ she is experiencing at the time.
Managing Troublesome Opioid-induced Respiratory Depression
Although different definitions of troublesome respiratory depression can be created, many factors, such as a patient’s condition and the health care setting, also will significantly influence how varying levels of respiratory depression might be perceived or managed. Opioid-induced respiratory depression can require emergent or urgent intervention. In the worst case scenario, opioid-induced respiratory depression results in respiratory or cardiorespiratory arrest. At times, it may be difficult to determine that excessive opioid effect is the cause of such an emergency, especially in the case of cardiorespiratory arrest. In particular, when hypoxic insult or injury results in activation of the sympathetic nervous system, auto resuscitation and post-arrest hyperventilation can mask the usual signs of the underlying pharmacological cause of the problem. Therefore, an appropriate index of suspicion for opioid-induced adverse events must be maintained in patients with acute pain requiring opioid analgesia. A patient who succumbs to excessive opioid effect will be narcotized, but such a patient may be arousable if strongly stimulated. Their pupils are likely to be pinpoint unless sympathetic activation has taken place. If the patient is apneic but has a pulse, an immediate call for assistance should be made, delivery of oxygen by bag and mask instituted, and naloxone, 0.2–0.4 mg administered, preferably intravenously. In severe opioid overdose cases, initial required doses of naloxone may total 1–2 mg or more, but such doses are not without potential complications
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(see below). If a patient is completely unresponsive and without a pulse, standard Cardiopulmonary Resuscitation (CPR) and Advanced Cardiac Life Support (ACLS) protocols should be immediately instituted and naloxone should be administered in any unclear cases of arrest in postoperative patients. If a narcotized patient is discovered, but responds to stimulation and is not completely apneic, a call for help should be made and an initial assessment to determine vital signs and oxygenation made. If oxygenation, pulse, and blood pressure are adequate, smaller doses (0.02– 0.04 mg) of naloxone can be administered and titrated for patient resuscitation. Naloxone was introduced into clinical practice in the late 1960s and numerous studies demonstrated its efficacy as an antagonist of opioidinduced respiratory depression. Reports of side effects (increases in heart rate and blood pressure) and more serious complications (pulmonary edema, cardiac arrest, ventricular tachycardia or fibrillation) soon followed [147,148]. Initial naloxone dose recommendations ranged from 0.4 to 0.8 mg. These high doses of naloxone contribute to exaggerated cardiovascular responses after drug administration and are rarely necessary after anesthesia or troublesome opioid-induced respiratory depression associated with postoperative analgesia. If intravenous access is not available, naloxone, in doses similar to those given IV, is effectively absorbed after intratracheal administration [149]. There are several causes for the increases in arterial blood pressure, heart rate and other significant hemodynamic alterations seen after naloxone reversal of opioids, including pain, rapid awakening, and sympathetic activation not necessarily due to pain. Nausea and vomiting increase when opioid-induced analgesia is antagonized with naloxone. Pain and awakening are not required for increases in heart rate and blood pressure after opioid reversal with naloxone [150]. Patients with coronary artery or cerebrovascular disease are more likely to be adversely affected by cardiovascular stimulation associated with naloxone reversal of excessive opioid effect. In addition, greater degrees of hypercapnia at the time of opioid antagonism will result in greater degrees of cardiovascular stimulation due to associated sympathetic stimulation. Metabolic stress is also more likely when patients receiving naloxone for opioid agonist reversal are hypothermic. Shivering can occur, and oxygen consumption and minute ventilation can increase two- to three-fold [151]. Alphaadrenergic agonists (e.g., clonidine) and opioids decrease transmission through the same preganglionic sympathetic neurons. This mechanism may explain clonidine’s effectiveness in blocking hemodynamic stimulation following naloxone reversal of fentanyl [152]. Onset of action of intravenous naloxone is rapid (1–2 min), and its half-life and duration of effect are short, approximately 30–60 min [153].
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Recurrence of respiratory depression after naloxone reversal of opioidinduced respiratory depression is due to the agent’s short half-life. Attempting to compensate for naloxone’s short duration of action by increasing the size of a single dose risks increasing the incidence and severity of unwanted side effects. Renarcotization occurs more frequently after the use of naloxone to reverse longer-acting opioids like morphine. In these circumstances, a continuous infusion of naloxone (e.g., 0.25–1.0 mg kg1 h1) may be helpful, but should not supplant good medical and nursing care and attentive observations of respiratory function. Preservation of analgesia following reversal of opioid-induced respiratory depression has long been a goal of researchers and clinicians. While numerous opioid antagonists have been studied in attempts to find the ideal compound (including nalorphine, levallorphan, naloxone and numerous others), none of these is optimal. However, Gan et al. [154] did report that a continuous infusion of naloxone (0.25 mg kg1 h1) combined with IV PCA morphine, not only reduced opioid-induced side effects, but actually decreased morphine requirements in 60 patients who had total abdominal hysterectomies. The authors suggest that low doses of naloxone may be optimal in that they can reverse opioid agonist-induced side effects and also produce some analgesia. Others have noted that low-dose naloxone produces analgesia [155,156]. Higher doses of naloxone (e.g., 41 mg kg1 h1) will lead to ineffective analgesia, while doses less than 1 mg kg1 h1 given to patients who receive intrathecal morphine, are more likely to spare analgesia and improve ventilation [157]. Nalbuphine has been extensively evaluated for this purpose [158,159]. Latasch et al. [160] reported that nalbuphine, 20 mg IV, adequately reversed respiratory depression but not analgesia in patients who had received fentanyl and nitrous oxide for general anesthesia. Zsigmond et al. [161] found nalbuphine, 0.1 mg kg1, did not elicit significant cardiovascular changes or pain in patients given fentanyl/nitrous oxide anesthesia for abdominal surgery. Unfortunately, many other investigators have documented the occurrence of significant pain, hypertension and tachycardia (often requiring pharmacologic intervention) following opioid reversal with nalbuphine [159,162]. Nalbuphine restores normal inspiratory neuronal activity after fentanyl in cats [99]. Restoration of spontaneous ventilation using small, titrated doses of nalbuphine (2.5 mg every 2–3 min) results in less pain than naloxone (0.08 mg titrated similarly) after fentanyl, isoflurane and nitrous oxide anesthesia [158]. Compared with naloxone, renarcotization is less likely after nalbuphine reversal of respiratory depression. Prophylactic administration of nalbuphine (200 mg kg1 bolus plus 50 mg kg 1 h 1 ) has been reported to prevent CO2 accumulation (PaCO2 4 50 mm Hg) and preserve analgesia from epidural morphine after thoracotomy [163]. Other studies, including evaluations of the use of
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nalbuphine with epidural hydromorphone [164] or nalmefene for IV PCA morphine [165] exist, but most of these examine control of more common opioid-induced side effects (e.g., pruritis) and not respiratory depression. Currently it would appear that low-dose (0.25–0.5 mg kg1 h1) IV naloxone infusions would be the best prophylaxis against opioid-induced side effects, including respiratory depression, no matter what the route of opioid agonist administration. VI.
Regional Analgesia
Systemic administration of opioid analgesics does not result in optimal pain control. There are no systemically administered drugs or drug combinations that produce good to excellent analgesia for severe pain and which block stress cascades or other deleterious responses and reflexes to noxious stimuli. In addition, systemic administration leads to generalized systemic drug action with associated side and adverse effects, all too often undesirable. Optimal analgesia, if it is also to potentially improve outcome, should be pre-emptive, consistent, continuous, and dynamic with minimal side or adverse effects. It is only with a multimodal approach to painful surgical or other conditions that such goals can be achieved, and regional analgesia has an integral role in such approaches [21]. In its broadest sense, regional analgesia includes any pain control modality that is targeted in some anatomically regional or local manner. Regional anesthesia or analgesia is frequently considered as peripheral (e.g., nerve or nerve plexus block) or central, i.e., neuraxial (e.g., intrathecal or epidural block). Many new adjuncts (e.g., opioids, alpha-2 agonists, ketamine; see below) are being developed and evaluated for use in regional analgesia. The most commonly used drugs for regional anesthesia and analgesia remain local anesthetics. Rare but very serious complications can occur after neuraxial anesthesia, including high spinal anesthesia, severe hypotension, cardiorespiratory arrest, nerve injury, and permanent spinal cord damage. Complications can also occur during peripheral regional anesthetic or analgesic techniques (e.g., significantly impaired respiratory function after brachial plexus block) [166,167]. In this chapter, we will only consider the usual and anticipated actions and effects of regional anesthesia and analgesia on respiratory function. A. Local Anesthetics
Regional anesthesia or analgesia with local anesthetics, in low concentrations, can contribute significantly to the goal of producing optimal pain control. When appropriately administered, they produce predictable and manageable side effects that usually are easily managed and
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can result in significant improvements besides excellent analgesia. Local anesthetics, if injected in large amounts or if unintentionally injected into the bloodstream, can lead to systemic toxicity. As the plasma concentration of a local anesthetic increases, symptoms range from circumoral tingling and numbness, lightheadedness, tinnitus and visual disturbances to muscle twitching, seizures, loss of consciousness, and finally coma and respiratory arrest. Cardiovascular depression can also occur. The risks of regional anesthetic techniques producing these complications are greatest at the time the block is performed. However, rarely, an epidural catheter can migrate either intrathecally or intravenously and if undetected, may result in exaggerated or unwanted effects. Low blood concentrations of local anesthetics minimally affect resting ventilation [168,169]. Some authors suggest that local anesthetics have no effect on ventilation during hypercarbia but a slightly stimulatory effect during hypoxia [169]. Others have found local anesthetics to have a stimulatory effect on ventilatory response to CO2. Gross et al. [170] found that lidocaine depressed the ventilatory response to CO2 transiently after a bolus intravenous injection while an infusion, after achieving steady-state levels, had a stimulatory effect on the hypercapnic ventilatory response. He suggested that lidocaine serum levels of 3.5 0.2 mg ml1 increased the slope of CO2 response curves and that ventilatory control in subjects receiving regional anesthesia must take into account the direct effect of local anesthetics on ventilation. Labaille et al. [171] showed a stimulatory effect of both IV infusion and lumbar epidural lidocaine. They studied the effects of intravenous and epidurally administered lidocaine on the control of ventilation in two groups of eight healthy unpremedicated subjects. In both groups, there was a significant increase in the ventilatory response to CO2, but resting minute ventilation and end-tidal CO2 values remained unchanged. The lidocaine serum levels were 3.14 0.82 mg ml1 in the intravenous group (bolus followed by an infusion) and 1.79 0.42 mg ml1 at 15 min and 2.22 0.47 mg ml1 25 min after the administration of 5 mg kg1 of lidocaine in the epidural group (upper levels T7–T10). These results suggest that the systemically mediated effects of local anesthetics given in the epidural space cause stimulation of ventilatory control mechanisms. Other studies (see below) address the ventilatory effects of spinal or epidural anesthesia per se but it appears that local anesthetic solutions administered for these blocks, when absorbed systemically, have minimal potential for causing respiratory dysfunction. B. Intravenous Regional (Bier) Block
Intravenous anesthesia for the upper or lower extremities can be provided by the sequence of: (1) physiologic exsanguination by the elevation and
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wrapping of the extremity; (2) application of a double tourniquet circumferentially around a portion of the extremity proximal to the surgical site; (3) inflation of the distal tourniquet to a pressure 50–100 mm Hg above the anticipated peak systolic blood pressure, and (4) venous injection of 0.5% lidocaine, 30–50 ml. The block lasts only as long as the tourniquet remains inflated. Premature tourniquet deflation, especially within minutes of injection of the local anesthetic, can lead to systemic toxicity, with concomitant respiratory compromise. There is no role for Bier blocks in the control of postoperative pain. C. Intra-articular Analgesia
Intra-articular injection of local anesthetic solutions prior to and after arthroscopic reconstruction of the anterior cruciate ligament can improve postoperative analgesia with no additional side effects [172]. The intraarticular administration of small doses of opioids has been widely studied, in particular after knee surgery, but results concerning analgesic efficacy have been inconsistent [173]. Some positive studies suggest that morphine is the opioid of choice for intra-articular opioid analgesia [174]. Since the doses usually employed are quite small (e.g., 1 mg morphine), there is no concern for respiratory depression after such intervention. Others have suggested that more lipid-soluble drugs such as sufentanil, 10 mg, are also effective in providing postoperative analgesia when administered intraarticularly [175]. D. Field blocks
Field block, or local anesthesia (e.g., incisional or wound infiltration with local anesthetic), is the simplest form of regional anesthesia. Easy to administer, and usually safe, field blocks have some appeal. For example, field blocks provide rapid-onset incisional analgesia. However, field blocks are not long lasting, do not lend themselves easily to redosing, and can carry some immediate risk of systemic toxicity if recommended limits on total drug dose are not adhered to and/or if intravascular injection occurs. Nevertheless, infiltration of painful superficial surgical sites is frequently practiced. Moiniche et al. [176] reported a systematic review of incisional local anesthesia for postoperative pain relief after abdominal operations. Of the more than 90 reports of infiltrative block, they found 34 studies concerning abdominal procedures, of which 26 met inclusion criteria for a total of 1211 patients. Operations included inguinal hernia repairs, hysterectomy, and cholecystectomy. There were no complications related to infiltrative block. However, with the exception of inguinal hernia repairs, there was no evidence for field block affecting postoperative pain.
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Brachial Plexus Blocks
Interscalene Approach
Shoulder and upper extremity pain, especially after surgery, can be severe enough to require hospital admission for patients who could otherwise be discharged from healthcare facilities. Blockade of the brachial plexus via the interscalene route can produce anesthesia for shoulder surgery and analgesia for pain control after shoulder surgery. Such analgesia can be extended for up to 24 h after single injection techniques or for days when administered via a properly inserted and secured catheter. Only regional analgesia can provide dynamic pain relief after shoulder surgery. The analgesia produced by brachial plexus block helps patients participate fully in physical therapy for chronic shoulder pain. An interscalene block is routinely placed at C6 level with the needle in the interscalene groove and directed caudally and posteriorly. About 30–45 ml of local anesthetic is injected. Due to the proximity of vagus, phrenic and cervical sympathetic nerves, a temporary block of these structures frequently takes place. Patients can experience mild dyspnea and/or hoarseness. Intravascular injection, especially if arterial, often causes immediate seizures and the usual associated respiratory problems. Pneumothorax can occur after interscalene block but is rare (0.2%) [177]. Rarely, epidural, subdural or intrathecal injection can occur and result in a high spinal with severe hypotension, loss of consciousness, and apnea. It is common for local anesthetic to spread to C3, C4, and C5 nerve roots during the interscalene block. Hence, phrenic nerve paresis causing hemidiaphragmatic paresis is very common. There is a consistent reduction in pulmonary function after pathological unilateral phrenic nerve paralysis [178] as well as during local anesthetic blockade of the phrenic nerve [179]. Urmey [180] studied the effects of unilateral hemidiaphragmatic paresis on routine pulmonary function and on chest wall motion during interscalene block. All patients except one had significant reductions in their FVC and FEV1. Peak inspiratory and expiratory flow rates decreased in five of eight patients. Interscalene brachial plexus anesthesia can reduce pulmonary function test results by 20–40 % [180,181]. Dyspnea can occur from hemidiaphragmatic paresis after interscalene block. Although it may be a distressing symptom, it is most often of minor consequence in normal healthy patients. The incidence of hemidiaphragmatic paresis can be as high as 100% [180]. Interscalene anesthesia can result in a decreased ability to breathe deeply, cough, and clear secretions. Normal and deep inspiration can result in increased ipsilateral abdominal and diaphragmatic paradoxical motion, possibly leading to regional atelectasis. Cardiac surgery can lead to left phrenic nerve paresis. Performance of a right interscalene block shortly after cardiac surgery has been reported to result in symptoms of respiratory distress [182].
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Interscalene block can cause hoarseness due to unilateral block of the recurrent laryngeal nerve. Occasionally recurrent laryngeal nerve paresis can be bilateral. In patients with preexisting vocal cord weakness or problems, the addition of interscalene-induced recurrent laryngeal nerve paresis can become a life-threatening problem. Specific history evaluating vocal cord function in patients undergoing this block is advisable. In patients with significantly compromised lung function, respiratory decompensation following interscalene block and associated bilateral recurrent laryngeal nerve paresis can occur. Interestingly, the addition of buprenorphine, 0.3 mg, to local anesthetic for brachial plexus block has been reported to increase the duration of postoperative analgesia in patients after upper extremity surgery from 5 to 17 h [183]. Clonidine too can improve analgesia when added to local anesthetic solution for brachial plexus block [184]. Axillary Approach
Compared with an interscalene approach, brachial plexus regional anesthesia via the axillary approach does not significantly reduce pulmonary function [185]. Except for intravascular injection and immediate systemic toxicity, most of the risks mentioned above which are associated with an interscalene block are significantly reduced with the axillary approach to the brachial plexus, because of the more peripheral nature of the block. Periclavicular Approach
Supraclavicular nerve blocks are less commonly performed. Nevertheless, because of the rapid onset of these nerve blocks, they are favored by some for rapid-onset regional anesthesia of the brachial plexus. Complete paralysis of the hemidiaphragm has been reported in 59% of patients receiving supraclavicular blocks [186], associated with decreases in pulmonary function. Patients who demonstrated only reduced diaphragmatic motion did not have significant impairment of pulmonary function. Unilateral phrenic nerve paralysis can also occur after the infraclavicular approach to brachial plexus block, resulting in hemidiaphragmatic paralysis and hypoxemia [187]. The risks of pneumothorax are increased with supra- or infraclavicular approaches to brachial plexus block. Unintended intravascular injection also can still occur. F.
Intercostal Nerve Blocks
Intercostal nerve blocks with local anesthetic can produce effective analgesia for related pain and can reduce opioid requirements, which may permit a more complete normalization of pulmonary function after thoracotomy,
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especially in high-risk patients [188–190]. Intercostal nerve blocks, achieved either by intermittent injections or with a continuous interpleural catheter, can be used for postoperative or posttraumatic pain management. Intercostal nerve blocks may help with pain control, enabling patients to reduce splinting, breathe deeply, and cough. However, intercostal nerve blocks can decrease FRC through paralysis of ribcage muscles [191]. Pneumothorax after intercostal nerve block is not rare and can be as high as 5.6% for each intercostal nerve blocked [192]. Ballantyne et al. [11] considered whether postoperative analgesia with intercostal nerve block versus systemic opioid analgesia produced benefits. Eleven relevant studies were found in their MEDLINE search from 1966 to 1995, but many problems existed with interpreting or including the studies. For example, most but not all studies evaluated one-time injections. Although several reports did find differences in surrogate measures of pulmonary function, overall statistical significance was not obtained. There was a suggestion that pulmonary complications might be reduced by intercostal nerve block if larger studies were performed. Compared with epidural analgesia (three studies), intercostal nerve block produced inferior analgesia but no significant differences in pulmonary function. The authors concluded that because of technical challenges with achieving persistent intercostal nerve block, it should not be instituted as first-line treatment. Nevertheless, the authors also found that intercostal nerve blocks may provide some benefit in reducing complications and that the block represents a potentially useful option when epidural treatment is not feasible. In an earlier meta-analysis, Kavanagh et al. [9] had also concluded that the value of single injections in producing intercostal nerve block was questionable, but that continuous intercostal nerve block through an indwelling catheter was probably beneficial. Cryoanalgesia has had some intermittent popularity for intercostal nerve blocks and the control of pain since it was first introduced by Lloyd [193]. It is achieved with application of a cryoprobe employing the Joule-Thomson effect, whereby carbon dioxide or nitrous oxide is released at high pressure and allowed to expand rapidly within the bulb of the cryoprobe [194]. This causes cooling of the probe tip to 50 to 70 C. When applied to peripheral nerves, localized freezing induces second-degree nerve lesions of axonotmesis [195]. Since the endoneurium remains intact, axonal regeneration takes place over weeks to months depending on the duration of exposure to the cryoprobe. Short- and longer-term analgesia after thoracotomy can be produced by cryoanalgesic techniques [196]; however, studies have suggested varying results and success [197]. Orr [198] and Pastor [199] have demonstrated superior analgesia and respiratory function after cryoanalgesia when compared with parenteral opioids. Moorjani et al. [200] reported on 200 thoracotomy patients who were randomized to either cryoanalgesia
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or conventional parenteral opioid analgesia. Pain scores were better and there was a trend toward better respiratory function (FEV1 and FVC) in the cryoanalgesia group. However, there was only one case of respiratory failure and death (parenteral group), limiting the power of the study. It is recognized that cryoanalgesia cannot provide complete analgesia after thoracotomy. Pain and inhibitory reflex signals transmitted via other nerves (e.g., vagus, sympathetic) are not affected by cryoanalgesia. Kavanaugh et al. [9] concluded that there appears to be little beneficial role for the routine use of cryoanalgesia after thoracic surgery. G. Interpleural Analgesia
Interpleural (sometimes called intrapleural) blocks involve the administration of local anesthetic between the two pleura. Typically, the surgeon inserts an epidural-type catheter at the end of surgery. The catheter is placed posteriorly, interpleurally, through the dermatome adjacent to the skin incision [201]. Local anesthetics such as bupivacaine (0.1–0.5%) are most often used and can be intermittently dosed or continuously infused. Advantages of interpleural analgesia when compared with thoracic epidural analgesia for postthoracotomy pain include ease of insertion. While it is potentially unsafe to insert a thoracic epidural catheter in patients who are anesthetized, interpleural catheter insertion in such patients is not problematic [202]. However, mixed results have been reported with the use of interpleural analgesia after thoracotomy [203,204]. Some authors report acceptable analgesia and/or improvements in lung function with this method of analgesia for pain, particularly after open cholecystectomy [205,206]. Few reports suggest that effective pain relief and/or equivalent improvements in pulmonary function after thoracotomy can be achieved with interpleural analgesia compared with thoracic epidural analgesia [202,207]. Others have not found interpleural techniques to produce either acceptable analgesia or significant improvements in lung function after thoracotomy. Reports of some benefit with this method note that analgesia and improvement in lung function are of relatively short duration [208]. Continuous interpleural analgesic technique may reduce pulmonary dysfunction after thoracotomy [209]. Removal of local anesthetic solution through chest tubes routinely placed after thoracotomy may be a reason for lack of efficacy of interpleural analgesia. A French review of 14 studies and a total of 318 thoracotomy patients receiving interpleural analgesia found four studies reporting no analgesia, five studies reporting modest, and five studies reporting good analgesia. Many of the studies had methodological flaws. Noted respiratory complications included pneumothorax, three patients with signs and symptoms of systemic toxicity (somnolence, disorientation,
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twitches), and one case of unilateral bronchospasm associated with injection of local anesthetic [210]. Although respiratory complications are rare with this technique, systemic levels of local anesthetic can approach toxicity [211]. Ballantyne’s review [11] evaluated seven papers comparing interpleural local anesthetic analgesia versus systemic opioid analgesia. No demonstrable difference in pulmonary function outcome was found in the meta-analysis. An earlier meta-analysis also could not report any definitive benefit with interpleural analgesia [212].
H. Paravertebral Block
Paravertebral block can be produced by placing an epidural-like catheter into the paravertebral space prior to chest closure [213]. It has been suggested that, when compared with thoracic epidural analgesia with local anesthetic, continuous paravertebral block (which can be similar to interpleural block) can produce superior static and dynamic pain scores [214]. Patients receiving paravertebral block also had better oxygenation (clinically not significant) and pulmonary function (PEFR). Markers of the stress response (plasma cortisol and glucose) were less in patients receiving paravertebral block. Side effects, especially hypotension, were found only in the epidural group. There was a trend toward reduced chronic pain at six months after surgery in the paravertebral block patients (3 versus 10). Total additional mean PCA morphine consumed at 48 h after surgery was less in the paravertebral group when compared with the epidural group (262 mg versus 210 mg). The authors concluded that paravertebral block was superior to epidural block after thoracotomy. The lack of opioid in the epidural infusion and the greater amount of bupivacaine (two times) administered in the paravertebral block group may have contributed to the observed differences. After cardiac surgery, paravertebral block is simpler to perform than thoracic epidural block and may provide equal analgesia and perhaps improvement in pulmonary rehabilitation after surgery [215]. I.
Neuraxial Blockade
Epidural Blockade with Local Anesthetics
The effects of epidural blockade with local anesthetics on respiratory function and pulmonary mechanics depend on the level and density of the block. Block height and spread is primarily determined by the level at which the catheter is placed and by the volume of solution injected or infused over time. Block density is largely determined by the concentration of local anesthetic injected. Conduction blockade may affect respiratory
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control through effects on chest wall receptors and respiratory muscles. In addition, as mentioned above, local anesthetics may affect the ventilatory drive through systemic effect after such drugs are absorbed into the bloodstream and delivered to the CNS via recirculation [170,171]. Occasionally, high spinal can complicate spinal or epidural anesthesia or analgesia and can result in respiratory arrest. The cause of respiratory arrest or apnea associated with such high spinals is the significant hypotension and subsequent brainstem ischemia that results. Spontaneous respiratory effort usually returns shortly after the restoration of blood pressure and circulation. This would not occur if the loss of respiratory effort was due to high spinal blockade of neuromuscular or brainstem function per se. Side effects of epidural local anesthetic include hypotension, motor weakness, urinary retention, and delayed ambulation but these problems can usually be avoided with low concentrations, e.g., 0.125% or less of bupivacaine. Catheter tip migration is rare, with an incidence of 0.1–0.2% [8], but rarely produces serious complications when drugs intended to be administered epidurally are given intrathecally. Patient-controlled epidural analgesia (PCEA) can bring benefits similar to those of PCA, e.g., improved analgesia and patient satisfaction. Most studies of the effects of epidural analgesia on respiratory function involve epidural anesthesia, with concentrations of local anesthetics greater than those used for postoperative analgesia. It is plausible that somatosensory and sympathetic blockade could produce deleterious effects through several mechanisms. Motor function of respiratory muscles could be weakened, spindle fiber feedback to the central controller could be disturbed; sensory feedback via the chest wall impaired, and sympathetic contribution to pulmonary blood flow and smooth muscle tone in the airways unfavorably altered. Coordination of the ribcage muscles could be impaired, resulting in abnormal contribution either to inspiration or to resting lung volumes [216]. Thus, conceivably, epidural blockade could lead to lower lung volumes, atelectasis, and hypoxemia. Several studies counter the above scenario with evidence that epidural anesthesia with local anesthetics has minimal effect on lung or respiratory function. FRC has been reported to undergo minor changes or to be stable in the presence of high thoracic epidural anesthesia (as long as significant hypotension was avoided) [217–219]. High epidural anesthesia increases FRC in healthy volunteers, perhaps resulting from caudad movement of the diaphragm (due to enhanced laxity in the abdominal muscles) and from a decrease in intrathoracic blood volume [216]. Ribcage expansion continues to contribute to tidal volume during high epidural anesthesia even when most of the ribcage muscles are paralyzed [216]. Even with cervical anesthetic block, changes in lung volumes (FEV1 and FVC) are modest (515%) and do not influence oxygenation or maximum inspiratory pressure
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[220]. Airway resistance has been documented to be minimally affected by thoracic epidural anesthesia [221] (Figure 13.10). Neither epidural nor spinal anesthesia appears to have significant impact on the ventilatory response to hypercapnia or hypoxia [222,223]. Several studies have shown that ventilatory response to hypoxia and hypercapnia is well maintained in lumbar or high cervico-thoracic epidural block in young or elderly patients [224,225].
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Figure 13.10 Forced expiratory volume in 1 sec (FEV1), vital capacity (VC) (percent predicted), and ratio of FEV1 over VC (percent) of 20 patients with chronic obstructive pulmonary disease or asthma in sitting (open bars) and in supine position before (gray bars) and during (black bars) epidural anesthesia with either ropivacaine (left, n ¼ 10) or bupivacaine (right, n ¼ 10). Mean SD; *P 5 0.05. VC and FEV1 decreased significantly both when attaining supine position and during high thoracic segmental epidural anesthesia. As a measure of airway obstruction, FEV1 as a percentage of VC increased significantly when attaining supine position and during segmental high thoracic epidural anesthesia (sTEA) (Data from Ref. 221).
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Epidural anesthesia also has minimal effect on gas exchange, even during one-lung anesthesia [226]. Other reports confirm that arterial blood gases are minimally affected by either spinal or epidural anesthesia [227–229]. Whatever alterations in ribcage motion that do occur are either minor or are adequately compensated for by other respiratory muscles [216]. In addition, as mentioned, systemic absorption of local anesthetic may actually mildly stimulate ventilation and counter any modest respiratory depression associated with neuraxial blockade. Some measures of lung function (inspiratory capacity, vital capacity, total lung capacity, and FEV1) may be modestly decreased during neuraxial block [219,230]. Although ERV can be decreased during motor blockade of the abdominal muscles, there is no change in FEV1 indicating no change in airway resistance. Intact abdominal muscle function and a normal PEFR can imply that cough mechanisms are maintained under epidural analgesia with local anesthetic. In patients with normal and diseased lungs, there are usually minimal decreases in tidal volume, minute volume and vital capacity with high thoracic levels of anesthesia, and the ability to cough is unlikely to be impaired. It is believed that effective pain relief with epidural local anesthetics, especially when combined with other drugs such as opioids, can improve postoperative respiratory function and reduce the incidence of postoperative respiratory problems [231]. Evidence supporting this hypothesis regarding pulmonary function will be reviewed here, while evidence concerning actual outcome data will be summarized below in a separate section. Compared with IM pentazocine, epidural infusions of lidocaine for postoperative pain relief in healthy patients after hip surgery did not result in significant ventilation–perfusion mismatch, shunt or hypoxemia [41]. Epidural analgesia using local anesthetics resulted in significantly fewer episodes of hypoxemia, central apnea, obstructive apnea and paradoxical breathing in patients while they were asleep and recovering from major surgery when compared with patients receiving parenteral opioids [24]. Effective pain control with thoracic epidural analgesia can partially restore vital capacity, FEV1, PEFR and FRC, all of which are significantly reduced in the postoperative period [48,232]. Dynamic tests of ventilatory capacity (e.g., PEFR, FVC, and FEV1), usually decreased after upper abdominal surgery, can be partially restored with good pain control [233]. Thoracic epidural block with local anesthetics provide good pain control and may partially reverse postoperative diaphragmatic dysfunction [51,234]. Manikian [51] demonstrated that thoracic epidural analgesia (0.5% bupivacaine) could at least partially reverse the pulmonary dysfunction associated with upper abdominal surgery. He reported 50% increases in tidal volume and abdominal contributions to breathing, increases in vital capacity, as well as decreases in respiratory rate.
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However, epidural analgesia with local anesthetics alone is not always sufficient and can produce significant side effects. Even with analgesic concentrations (e.g., 0.125% bupivacaine), significant hypotension can result from thoracic epidural analgesia [235,236]. Supplementation with NSAIDs does not always render analgesia effective [237]. Other drugs, most frequently opioids, are combined with local anesthetics in an attempt to provide complete analgesia. Epidural Opioids
The administration of opioids via the epidural route has advantages, since opioids produce both pre- and postsynaptic analgesic effects in the dorsal horn of the spinal cord. Opioid action includes modulation of nociceptive input and, unlike local anesthetics, epidural opioids do not cause sympathetic or motor block. Neuraxial opioids also produce only minor degrees of sensory block except analgesia. The lack of motor block should prevent epidural opioids from altering respiratory function through mechanical effects. However, in evaluating most studies of epidural opioids alone, when compared with IV opioids, there may be little advantage to an analgesic strategy that relies on epidural versus IV opioids [8]. Neuraxial opioids do not consistently produce dynamic analgesia. It appears that combinations of epidurally administered opioid and local anesthetic permit optimal analgesia while minimizing side or adverse effects of either opioids (e.g., respiratory depression, pruritis, nausea) or of local anesthetics (motor block, hypotension) [8]. Concentrations and total hourly drug doses can be significantly reduced when the drugs are combined. There is little information using isobolographic analyses to determine if combining opioids and local anesthetics results in synergism, nor to explain the efficacy of the drug combination. Nevertheless, combining opioids and local anesthetics is popular and does produce dynamic analgesia most consistently [8,238]. It has been suggested that optimal drug concentrations for bupivacaine and fentanyl are 0.125% and 4 mg ml1, respectively [239]. Respiratory depression can occur after epidural opioid administration. Biphasic respiratory depression following epidural morphine was reported by Kafer et al. in 1983 [240]. Seven chronic pain patients were evaluated for the effects of 0.1 mg kg1 of lumbar epidural morphine. Maximal depression of the slope of the ventilatory response to CO2 occurred at 1–2 h post-injection with recovery of ventilatory drive at 4 h. Recurrence of respiratory depression was manifested at 8 h but not 12 or 24 h after injection of epidural morphine. The authors postulated two separate mechanisms producing the biphasic depression. Early depression was due to systemic absorption of drug, while the delayed depression was due to rostral spread of drug within the neuraxis [241]. Delayed ascension of morphine over 4–8 h
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within the neuraxis was associated with other signs (e.g., analgesia) of rising segmental neuraxial opioid effect, and was documented by Bromage et al. in 1982 [258]. Camporesi et al. [242] also reported the ventilatory effects of 10 mg of epidural versus IV morphine (in 10 ml) in 10 human volunteers. Maximal respiratory depression after IV morphine was at 0.5 h after injection but was delayed until 6–10 h after epidural injection. Significantly greater respiratory depression was evident after epidural versus IV morphine. On the other hand, Sandler et al. [243] and others have demonstrated that lipophilic opioids such as fentanyl produce equivalent blood drug levels whether the drugs are administered intravenously or epidurally. This has led some to conclude that there is no advantage to epidural administration of lipophilic opioids, while others contend that regional spinal effects are still superior after epidural administration of lipid-soluble opioids. Limited cephalad migration of lipid-soluble opioids such as fentanyl has been thought to confer some protection against respiratory depression [120]. Lipid-soluble opioids administered in the epidural space are less likely to cause biphasic respiratory depression when compared with morphine [124]. Epidural fentanyl administered at the lumbar level produces maximum cervical CSF fentanyl concentrations that can be 10% of the CSF concentration in the lumbar CSF [244]. In general, lipid-soluble opioids are recommended when epidural catheter tip location is at or near the dermatomal sites of pain, and less lipid-soluble opioids (e.g., morphine) are recommended when the catheter tip is more distant from the site of nociception. Successful dynamic analgesia is more likely when the epidural catheter tip is at or near the site of pain (e.g., thoracic versus lumbar) [8]. Clinically significant depression can occur after either epidural bolus or infusion of fentanyl. The ventilatory response to CO2 is depressed after 200 mg of epidural fentanyl [245]. Severe respiratory depression and arrest have been reported after 100 mg boluses of epidural fentanyl [120]. Epidural fentanyl infusions of 0.5–1.0 mg kg1 h1 can depress the ventilatory response to CO2 [246]. Overall, in 29 studies involving over 600 patients receiving epidural fentanyl, the incidence of troublesome respiratory depression with epidural fentanyl was 1.8% [120]. Hansdottir et al. [247] studied the effects of continuous epidural sufentanil and bupivacaine infusion after thoracotomy in 37 patients. Patients received sufentanil at either the thoracic or the lumbar level at a rate of 4–20 mg h1 (1 mg ml1). A third group received sufentanil plus bupivacaine at the thoracic level. Lung volumes (e.g., vital capacity) were reduced in all three groups by approximately 50%. There was a trend toward better responses (minute ventilation, inspiratory flow and occlusion pressure) to 7% CO2 in patients receiving thoracic epidural sufentanil plus bupivacaine. Sedation and hypercapnia occurred most frequently in the lumbar epidural sufentanil group. Twenty-four hours after discontinuation
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of the infusions, there was complete recovery of CO2, responses only in patients receiving thoracic epidural sufentanil plus bupivacaine. The authors concluded that after thoracotomy, epidural sufentanil analgesia is optimal when tailored to the site of nociceptive input and combined with bupivacaine [247]. The true incidence of troublesome respiratory depression related to neuraxial opioid administration, whether epidural or intrathecal, is not known. Many factors, such as differences in patient populations and surgery, types of drugs and drug delivery methods, adjuvant and other medications concomitantly administered, and the level of monitoring, can all influence the real and/or perceived magnitude and duration of respiratory depression associated with neuraxial opioids. Although neuraxial opioids have been suggested to carry a reduced risk of troublesome respiratory depression when compared with systemically administered opioids [248], the incidence of troublesome opioid-induced respiratory depression is likely to be similar regardless of the route of drug administration. The reported incidence of respiratory depression after epidural opioid administration ranges from 0.1% to 1% [8,248,249], similar to that of opioids given by other routes. The onset of troublesome respiratory depression associated with epidural opioid drugs is usually slow to develop. Based on what is known about CSF circulation, peak ventricular morphine concentrations can be expected to develop within 2–6 h following lumbar epidural morphine injection [250,251]. Lipid-soluble opioids are removed from the epidural space more rapidly than the hydrophilic molecule morphine. Rapid vascular absorption of lipid-soluble opioids limits drug availability for rostral migration within the neuraxial region but can deliver opioid to the brain via systemic circulation. After epidural injection, significant blood concentrations of fentanyl and sufentanil can be measured within 5–10 min or less [244,252]. Thus, after epidural injection, peak CSF levels of fentanyl or sufentanil occur in less than 20 min, whereas peak levels of morphine can take up to 4 h to develop. Early- as well as late-onset respiratory depression has been reported with most opioids after epidural injection, including fentanyl, meperidine, hydromorphone, and sufentanil [253–255]. With higher doses of drugs such as sufentanil, direct spread within the CSF and/or via epidural veins cannot be ruled out as a possible mechanism of associated troublesome respiratory depression. Epidural administration of very lipid-soluble opioids such as sufentanil may not be appropriate. The dose of sufentanil, when administered via the epidural route, may need to be larger than that of sufentanil given IV in order to produce the same level of analgesia. It may be that sufentanil, because of its extreme lipophilicity, is excessively absorbed by epidural fat, reducing effective neuraxial action.
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Keys to minimizing clinically significant respiratory depression associated with epidural opioid administration are: (1) supervising, managing and monitoring such pain strategies with a well-organized acute pain service [8]; (2) regularly monitoring, at least once every hour, respiratory rate and level of consciousness during therapy; (3) maintaining monitoring for an adequate period of time (e.g., up 12 h after last epidural bolus of morphine), and (4) administering supplemental oxygen. Intrathecal Anesthesia with Local Anesthetics
With the exception of intrathecal opioid administration, intrathecal analgesia is not commonly used for postoperative pain control. In the immediate postoperative period, spinal block from intraoperative anesthesia can persist. Ventilatory mechanics are minimally affected by intrathecal (i.e., subarachnoid or spinal) anesthesia with local anesthetic. The effects of intrathecal anesthesia are different on inspiratory and expiratory function because of the muscle groups involved. Inspiratory function is minimally affected as the principal inspiratory muscle, the diaphragm, is spared even with high levels of thoracic blocks because its innervation derives from the cervical plexus. In addition, the accessory muscles of inspiration (e.g., scalenes and sternocleidomastoids) are not affected because of their craniocervical innervation. Diaphragmatic function is also more efficient due to relaxation of abdominal muscle tone. Any existing blockade of other inspiratory muscles, like external intercostals, is therefore well compensated. Expiratory function can be affected because expiratory muscles (e.g., internal intercostals, abdominal wall muscles) are frequently weakened by spinal anesthesia. Although at-rest expiration is mostly passive, active effort is required during coughing. Thoracic levels of spinal anesthesia are associated with significant reductions in maximal expiratory pressures and flow rates [64]. Spinal anesthesia is not associated with statistically significant changes in the resting ventilation or the response to the single-breath CO2 test [223]. Intrathecal anesthesia with bupivacaine can lead to an increased responsiveness to hypercapnia [64], which may reflect the relatively small contribution of peripheral chemoreceptors in a rebreathing hypercapnic ventilatory response test. A single-breath CO2 test predominantly affects the peripheral chemoreceptors as opposed to a CO2 rebreathing technique where central chemoreceptors are involved. An increased response to CO2 after intrathecal block could be due to blockade of inhibitory chest wall afferents. The response to isocapnic hypoxia is well preserved in lumbar as well as thoracic levels of anesthesia [256]. Similarly, the response to hypercapnic hypoxia is also not affected by age or anesthetic level [225]. Recently, the
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effects of intrathecal bupivacaine and sameridine (a new compound with both local anesthetic and opioid properties) were studied [257]. Neither drug significantly affected the hypoxic or hypercapnic ventilatory response in healthy volunteers, nor did either drug affect the resting end-tidal CO2 although bupivacaine decreased the tidal volume with a compensatory increased respiratory rate. Sameridine depressed the hypercapnic ventilatory response to a small degree.
Intrathecal Opioids
The intrathecal administration of opioids has several advantages. It is a relatively simple and reliable technique; because there is no dura for drug to penetrate and because drug is deposited very close to the intended site of action, very low doses are effective. In addition, intrathecal administration of morphine, because of its hydrophilic nature and retention within the CSF, produces analgesia for up to 24 h [258]. In comparison, epidural doses for morphine must be approximately 10 times greater to produce similar analgesia. Although initial doses when intrathecal morphine was first introduced ranged from 1.0 to 5.0 mg or more, currently doses range from 0.1 to 1.0 mg with 0.2–0.4 being the most commonly used. Doses of 10–25 mg of fentanyl and 2.5–10 mg of sufentanil are also used [258]; meperidine has also been employed. The major limiting factor in the continuous administration of intrathecal opioids for acute pain is the lack of a catheter delivery system that allows delivery of drug with minimal technical problems or complications. Respiratory depression after intrathecal opioids has been well studied [248]. Bailey et al. [258] studied 20 male volunteers for 24 h after intrathecal doses of placebo and morphine doses of 0.2, 0.4, or 0.6 mg. Dose-related analgesia and respiratory depression were documented and significant effects lasted up to 20 h. For example, after 0.6 mg of intrathecal morphine, the slope of the ventilatory response to CO2 was 550% at 19.5 h after drug injection (Figure 13.11); this is consistent with the slow removal of morphine from the CSF. Respiratory rate did not correlate with drug dose. Oxyhemoglobin desaturations (SpO2 5 90%) occurred in all groups but its incidence was dose related. Supplemental oxygen was consistently effective in restoring normoxia. Arterial CO2 tensions were also dose related, with a first peak at 6.5–7.5 h, consistent with what is known about CSF circulation [253]. Secondary peaks were noted and were related to a time effect (placebo group) and possibly to sleep. The highest PaCO2 noted was 55.8 mm Hg in a subject receiving 0.6 mg of intrathecal morphine. The effect of intrathecal (IT) morphine (MS) on the ventilatory response to hypoxia was also studied by Bailey et al. in human volunteers
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Mean Slope of VE vs CO2 (L · min−1· mm Hg−1)
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Figure 13.11 Mean ventilatory response to carbon dioxide vs. time before (time 0) and hours after intrathecal morphine sulfate by dosage group: 0, 0.2, 0.4, or 0.6 mg (Data from Ref. 258).
[101]. Thirty subjects were randomly assigned to receive one of three treatments in a double-blind study: (1) IV and IT sterile saline; (2) IV MS (0.14 mg kg1) and IT saline; or (3) IV saline and IT MS (0.3 mg). This study design not only allowed a determination of the effect of intrathecal morphine on the ventilatory response to hypoxia, but it also allowed distinction between peripherally and centrally mediated effects. This was because intrathecal morphine produces minimal to no blood morphine or morphine metabolite levels and therefore would not be expected to exert an effect at the peripheral chemoreceptor. If opioidmediated depression of the acute ventilatory response to hypoxia was via drug action at the peripheral chemoreceptor, then intrathecal morphine should not depress the ventilatory response to hypoxia. The authors found that both IV MS and IT MS depressed the acute ventilatory response to hypoxia, but differently. IV MS depression of the acute ventilatory response to hypoxia was maximum at 3.5 h, compared with 7 h after IT MS. In addition, 12 h after injection, significant depression of the acute ventilatory response to hypoxia was still evident only in those subjects receiving IT MS. This study also determined that opioid-induced depression of acute ventilatory response to hypoxia is sufficiently explained by central
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Figure 13.12 Time course of the ventilatory response to acute hypoxia in three different groups of human study subjects who were healthy volunteers: (1) intravenous (IV) and intrathecal (IT) sterile saline; (2) IV morphine (MS) (0.14 mg kg1) and IT saline; or (3) IV saline and IT MS (0.3 mg). Values are means SE (Data from Ref. 101).
mechanisms (Figure 13.12). It is known that opioids also can depress peripheral chemoreception of hypoxia and ventilatory drive. The intrathecal administration of lipid-soluble opioids also produces intense analgesia but because of vascular absorption, the duration of action is shorter, lasting 1–3 h. Respiratory depression can occur quite soon after intrathecal injection of fentanyl or sufentanil. Respiratory arrest has been reported after intrathecal sufentanil, 10 mg, used in a combined spinal–epidural technique (CSE) for labor analgesia [259]. Lu et al. studied the effects of intrathecal sufentanil (12.5, 25, and 50 mg) on the ventilatory response to CO2, in 18 healthy female volunteers [260]. Serum sufentanil levels correlated with intrathecal sufentanil doses as well as with respiratory depression. Analgesia for experimental lower extremity pain was similar in onset and magnitude for all doses. The data are in keeping with the
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understanding that lipid-soluble opioids provide effective neuraxially mediated analgesia at low doses but produce systemically mediated effects at higher doses [261,262]. More recent animal data suggest that more than one explanation underlies the differences in pharmacokinetcs of various opioids after intrathecal injection. Morphine does have slow clearance into plasma but also has a low spinal cord volume of distribution. Fentanyl distributes rapidly into the epidural space and fat, and sufentanil has a high spinal cord volume of distribution [263]. The authors determined a rank order for meningeal permeability with alfentanil 4 fentanyl 4 morphine 4 sufentanil. The authors believe that hydrophobicity (octanol:buffer distribution coefficient) determines the ranking with drugs of intermediate hydrophobicity (alfentanil ¼ 129) being more permeable than drugs of greater (fentanyl ¼ 955, sufentanil ¼ 1,737) or lesser (morphine ¼ 1) hydrophobicity. Thus, known potency ratios for intravenous dosing of the various opioids may not hold true for injection of these drugs into other compartments such as the subarachnoid space [263]. The discussion of the possible nuances of CSF pharmacokinetics is beyond the scope of this chapter and indeed, significant difficulties in modeling the CSF pharmacologically have been described [264]. In the clinical setting and after surgery, intrathecal morphine can produce troublesome respiratory depression with doses as low as 0.3 mg [265]. Currently, unless patients are monitored in an intensive care unit or similar setting, intrathecal morphine doses most commonly used and presumed to be quite safe are 0.2–0.3 mg. Indeed, in some settings (e.g., after Cesarean section), it has been suggested that there is no advantage to giving patients more than 0.1 mg of intrathecal morphine [266]. A report of over 5,000 surgical patients treated with intrathecal morphine documented an incidence of respiratory depression (respiratory rate 5 8 or PaCO2 4 50 mm Hg) of 3% that was always detected by nursing observation, never life threatening, and always responsive to treatment with naloxone [267]. The authors reported no deaths or other serious complications related to intrathecal morphine analgesia. The typical intrathecal morphine dose ranged from 0.2 mg (TURP, vaginal hysterectomy) to 0.4–0.5 mg (hip, knee, lower abdominal surgery), and to 0.6–0.8 mg (upper abdominal, thoracic surgery). Patients were not sent to an ICU merely to monitor the effects of intrathecal morphine, but many of the patients treated with the higher doses of intrathecal morphine (0.5–0.8) did receive intensive care. All patients received mandatory hourly evaluations for the first 24 h. Other studies confirm that with proper selection of patient candidates and with appropriate postoperative surveillance and care, doses of 0.2–0.4 mg of IT MS can be safely used on regular hospital wards. Clinical studies continue to appear supporting the efficacy and safety of this dosage range [268]. Since the onset of intrathecal morphine takes several hours
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to peak, combinations of morphine with a more rapid-acting (i.e., lipidsoluble) opioid (e.g., fentanyl, sufentanil) may provide for immediate onset and long-acting analgesia. Combinations of sufentanil and morphine also can provide superior analgesia when compared with IV PCA morphine in patients for the first 24 h after thoracotomy. However, after the first postoperative day, no further benefit is produced and spirometric data in patients who were treated with intrathecal opioids is no better than in those patients only receiving PCA morphine [269]. VII.
Other Analgesic Agents
In the past few decades, significant achievements have been made in the understanding of spinal nociception and analgesia. Numerous receptor types involving opioids, serotonin, muscarinic agents, adenosine, gammaaminobutyric acid, glycine, somatostatin, and substance P have been identified as having potential roles in nociception. Some of these alternative analgesics will be discussed below. A. Agonist/Antagonist Compounds
Partial agonists or mixed agonist/antagonist compounds were introduced into clinical practice decades ago with the purported advantages of superior safety profiles and reduced addiction potential. Currently there are several agonist–antagonist opioids in use in the United States, including pentazocine, butorphanol, nalbuphine, dezocine, and buprenorphine. There is typically a ceiling to the effect of these agents and indeed this does limit the occurrence of clinically significant respiratory depression. Dysphoria is more common with these agents. This class of drugs has a limited role in the treatment of acute pain, especially if the pain is moderate to severe. These agents may play a role in the control of opioidinduced side effects and excessive respiratory depression. Buprenorphine is a partial agonist at the mu receptor. The other compounds are mu antagonists and full or partial agonists at the kappa receptors. Buprenorphine has an especially high affinity at the mu receptor while its actions at kappa receptors are probably minimal. Agonist–antagonist opioids are less prone (but not immune) to abuse because they cause less euphoria, and they are associated with less drugseeking behavior and physical dependence. Pentazocine is one-half to one-fourth as potent as morphine. Moderate analgesia results from 10 to 30 mg IV or 50 mg orally, and ceilings to both analgesia and respiratory depression occur after 30–70 mg of pentazocine. Although the potential for abuse is less than with morphine, prolonged use of pentazocine can lead to physical dependence. The drug is
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not particularly useful in reversing the respiratory depressant effects of fentanyl [270], but can precipitate opioid withdrawal in addicts [271]. Pentazocine has been reported to have unfavorable respiratory effects when used for postoperative analgesia [41]. Healthy patients who received pentazocine for postoperative pain relief had increased venous admixture, due to both increases in ventilation–perfusion mismatch and true shunt. This pattern of poor pulmonary function persisted for at least three days and caused significant hypoxemia for the first three days after surgery. Interestingly, patients receiving pentazocine, but not epidural analgesia with local anesthetic, demonstrated elevated diaphragms. Buprenorphine is a thebaine derivative, mu receptor partial agonist, and similar in structure to morphine, but approximately 33 times more potent. It also binds to delta and kappa receptors but activity at the latter two sites is relatively insignificant. Although buprenorphine is highly lipophilic, its opiate receptor association and dissociation is slow. Whereas fentanyl dissociates rapidly from mu receptors (T1/2 ¼ 6.8 min), buprenorphine has a higher affinity and takes much longer (T1/2 ¼ 166 min); thus, plasma levels do not parallel central nervous system effects [272]. Buprenorphine’s onset of action is slow and its peak effect may not occur for 3 h. Its duration of effect is prolonged (up to 10 h). Buprenorphine produces respiratory depression that may be significant, with a ceiling after 0.15–1.2 mg in adults. Higher doses do not produce further respiratory depression and may actually result in increased ventilation (predominance of antagonistic actions) [273,274]. Reversal with naloxone is limited due to buprenorphine’s high affinity for and slow dissociation from the mu opiate receptor. The addition of buprenorphine, 0.3 mg, to a local anesthetic solution used for brachial block has been reported to markedly extend postoperative analgesia without adverse respiratory effects [183]. Nalbuphine binds to mu as well as kappa and delta receptors. Nalbuphine acts as an antagonist at the mu receptor and an agonist at the kappa receptor. Activation of supraspinal and spinal kappa receptors results in limited analgesia, respiratory depression and sedation [275]. Although 10 mg of nalbuphine produces similar sedation, analgesia and respiratory depression as 10 mg of morphine, this equivalency does not persist at higher doses. Maximal analgesia occurs after approximately 30 mg of nalbuphine in a 70 kg patient. Nalbuphine also causes other typical opioid side effects. The drug is only available for parenteral use. Onset of effect is rapid (5–10 min) and duration long (3–6 h) due to a long plasma elimination half-life (5 h). B. Tramadol
Tramadol is a synthetic 4-phenyl-piperidine analogue of codeine with a dual mechanism of action [276]. Tramadol stimulates mu, and possibly to
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a lesser extent delta and kappa opioid receptors [277], and like tricyclic antidepressants, it also activates spinal inhibition of pain by decreasing the reuptake of norepinephrine and serotonin. Tramadol is prepared as a racemic mixture, and the enantiomers of tramadol have complementary and synergistic antinociceptive effects [278]. Thus, tramadol represents a drug with multiple mechanisms of analgesic action. The O-demethylation metabolite of tramadol shows higher affinity for the mu opioid receptor than its parent compound [279]. Alpha-2 agonist inhibitors can reduce the antinociceptive effect of tramadol. Tramadol is unlikely to produce troublesome respiratory depression at recommended doses but can at greater doses. Tramadol is one-fifth to one-tenth as potent as morphine and can be given by numerous parenteral routes. Recommended doses range from 50 to 100 mg every 4–6 h with a maximum daily dose of 400 mg. The effects of tramadol on the ventilatory response to CO2 have been studied in human volunteers [280]. On average, tramadol produced a 30% reduction in the sensitivity to CO2 but variability was significant. The authors suggested that tramadol depresses brainstem integrating respiratory centers via drug action at mu opioid receptors. Since tramadol did not affect oxygen consumption or the bispectral index, the authors concluded that metabolic or arousal state effects did not contribute to the respiratory effects of tramadol. Some other investigators have reported results that conflict with these [281]; however, the methods used have been called into question [280]. Others have also reported dose-dependent depression of the ventilatory response to CO2 after tramadol in humans. Tramadol has been reported not to depress the ventilatory response to hypoxia [282]. The respiratory depressant effects of tramadol in humans have been reported to be incompletely (50–70%) reversed by naloxone, suggesting that other mechanisms besides mu opioid receptor-mediated actions may play a modest role in the respiratory effects of tramadol [280]. Naloxone has been reported to completely reverse the effects of tramadol on ventilatory control in animal studies [283]. Intravenous tramadol has been reported to be as effective as epidural morphine and superior to IV morphine after thoracotomy [278]. Others report no significant differences between morphine and tramadol for postoperative pain management [284]. Epidural tramadol has been reported to be as effective as bupivacaine in the management of postoperative pain in children [285]. The addition of tramadol to local anesthetic for brachial plexus block prolongs sensory and motor block by 1–2 h without side effects [286]. It has been suggested that, due to systemic absorption, the epidural administration of tramadol has no advantage over intravenous administration [287].
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Alpha-2 adrenergic receptors are located on primary afferent terminals on neurons in the superficial laminae of the spinal cord and within several brainstem nuclei implicated in analgesia [288]. Several lines of evidence now exist confirming that the alpha-2 agonists produce not only spinally mediated analgesia, but other effects as well (Figure 13.13). Alpha-2 adrenergic agonists were introduced three decades ago as antihypertensive agents, but their acceptance by patients was limited due to
Physiology of Alpha-2 Adrenoceptors
Sedation
Bradycardia Decrease Tachycardia
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Figure 13.13 Responses that can be mediated by a2-adrenergic receptors. The site for the sedative action is in the locus coeruleus of the brain stem, whereas the principal site for the analgesic action is probably in the spinal cord; however, there is clear evidence for both a peripheral and a supraspinal site of action. In the heart, the dominant action of a2 agonists is a decrease in tachycardia (through block of the cardio-accelerator nerve) and bradycardia (through a vagomimetic action). In the peripheral vasculature, there are both vasodilatory action via sympatholysis and vasoconstriction mediated through the receptors in the smooth muscle cells. The mechanisms for the antishivering and diuretic actions have yet to be established firmly (Data from Ref. 289).
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accompanying sedation. Alpha-2 adrenergic agonists produce their clinical effects through alpha-2 receptors, of which there are at least three subtypes [289]. Their actions are not mediated through opioid receptor mechanisms [290]. The alpha-2A subtype appears to be responsible for anesthetic responses, including sedation and analgesia [289]. To date there are no subtype selective agonists to allow separation of desired effects (e.g., analgesia) from unwanted effects (e.g., hypotension) mediated by other alpha-2 subtype receptors. The alpha-2 agonists produce a spectrum of effects including analgesia, anxiolysis, sedation, and sympatholysis. Although the alpha-2 adrenergic agonists can produce profound hypoxemia in some species [291], they do not appear to have significant respiratory depressant effects in humans, even after large doses [288]. Some of the discrepancies found concerning the respiratory effects of alpha-2 agonists in humans might be related to type-one errors, to inadequate control groups or baseline data for comparison. In addition, confounding factors such as the unmasking of opioid-induced respiratory depression once pain is relieved by an alpha-2 agonist may also play a role [288]. Finally, sedation from alpha-2 agonists may result in some decrease in resting ventilation that is more related to decreases in wakefulness than direct depression of respiratory centers. Bailey et al. [292] reported the effects of clonidine (0.3–0.4 mg orally) alone or in combination with intravenous morphine in human volunteers. Clonidine alone did not depress either the ventilatory or the occlusion pressure response to CO2 rebreathing (Figure 13.14), but did shift these responses to the right (Figure 13.15). Clonidine also did not potentiate the effect of morphine on these parameters. Oxyhemoglobin desaturations were statistically significant but clinically minor after clonidine. Jarvis et al. [293] had similar findings to Bailey et al. [292] after evaluating clonidine and alfentanil in human volunteers. They found clinically minor decreases in minute ventilation and inspiratory flow attributable to clonidine alone and no potentiation of alfentanil-induced respiratory depression by clonidine. Belleville et al. [294] rapidly (over 2 min) administered dexmedetomidine (up to 2 mg kg1) to human volunteers. Although no volunteer reportedly experienced arterial oxyhemoglobin desaturations less than 90%, irregular breathing and apnea were noted. There was a reduction in tidal volume with a concomitant reduction in minute ventilation, and the ventilatory response to CO2 was shifted to the right and depressed (Figure 13.16 and Figure 13.17). The respiratory depressant effects of these rapidly administered doses of dexmedetomidine could be central as there are alpha-2 adrenergic receptors in the brainstem [295]. Ooi et al. [296] also found decreases in the slope of the ventilatory response to CO2 that ranged from 20% to 44% after intravenous clonidine. Intravenous dexmedetomidine, administered slowly as now recommended, does not appear to depress respiratory rate or ventilation
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Slope VE/CO2 (l· min-1· mm Hg-1)
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Figure 13.14 Slope of the ventilatory response to CO2 (mean SEM; l min1 mm Hg1) over time (time 0 ¼ control) for three drug groups: C ¼ clonidine nidine (0.3–0.4 mg orally), M ¼ morphine, 0.21 mg kg1 IM), or both drugs in the same doses combined. Changes were statistically significant for the morphine and morphine plus clonidine groups but not for the clonidine group (see text) (Data from Ref. 292).
(as reflected in arterial PCO2) and may even improve oxygenation in critically ill patients [297]. Hypotension and bradycardia occur frequently upon initiating dexmedetomidine infusions, especially when they are rapid or in patients with cardiac problems [297]. The effects of epidural alpha-2 agonists are of significant interest, as it appears that this route may be best for taking advantage of this class of drugs’ mechanism of action at the spinal level. Penon reported, in seven human volunteers, that epidural clonidine did not affect resting ventilation but did depress the slope of the ventilatory response to CO2 by approximately 33% [298]. Others have reported ventilatory rhythm disturbances and or upper airway obstructive episodes after clonidine orally [299] or epidurally [300] and after dexmedetomidine [294]. Eisenach et al. [301] found that 700 mg of epidural clonidine produced significant systemic levels of clonidine but only small (4 mm Hg) increases in PaCO2. The effects of the alpha-2 agonists on obstruction and the ventilatory response to hypoxemia merit further study. Narchi et al. [300] did report oxyhemoglobin desaturation in three of six patients who received epidural clonidine.
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35 C M C+M
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Figure 13.15 VE50 (mean SEM; l min1) over time (time 0 ¼ control) for the same three drug groups in Figure 13.14. All three groups had significant changes; statistically greater depression occurred in the morphine and morphine plus clonidine groups when compared with the clonidine group (see text) (Data from Ref. 292).
Alpha-2 agonist use as a substitute for opioids can reduce most opioidrelated side effects. They can also reduce opioid requirements by more than 50% and help control the hyperdynamic state after severe injuries such as burns [302]. Clonidine premedication can reduce the need for opioid analgesia and thus may provide an analgesic approach that produces less respiratory depression or risk. Clonidine, when combined with local anesthetics, can prolong sensory blockade produced by local anesthetics and can reduce the amount of local anesthetic required to produce postoperative analgesia [303]. Bier block with clonidine has been reported to alleviate sympathetically maintained pain [304]. The addition of clonidine to brachial plexus block also prolongs analgesia and/or enhances the quality of block without respiratory problems [184]. Alpha-2 agonists have been used successfully for postoperative pain control in diverse surgical patient populations and via many routes. As with opioids and other agents (e.g., neostigmine), the intra-articular administration of clonidine has been reported to produce clinical analgesia [305]. Epidural clonidine can improve analgesia without adding to the side effect profile when combined with local anesthetics for postoperative pain [306].
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Figure 13.16 Average minute ventilation while breathing room air for all dose groups. For clarity, SEM is shown for the placebo and 0.5 and 1.0 mg kg1 groups. See Figure 13.1 for explanation of the measurement times. P 5 0.05 different from the baseline (time ¼ 0) measurements within a dose group. For the 1.0 mg kg1 group, a significant difference from baseline was found by ANOVA, but a specific time could not be isolated at the P 5 0.05 level (Data from Ref. 294).
Small doses of intrathecal clonidine (30 mg) are as effective as 2.5 mg of sufentanil for labor analgesia in parturients [307]. Intrathecal clonidine does not produce hypotension and bradycardia to the degree that can be seen after intravenous administration [308], but hypotension can occur as the dose of clonidine is increased. Although the above-quoted studies and other reports substantiate that centrally administered clonidine alone can produce postoperative analgesia with minimal respiratory depression [309], clinical success is not always uniform or consistent. Thus, the ability of clonidine or of this class of drug to produce analgesia alone after intrathecal injection continues to be debated [310]. In summary, the alpha-2 agonists seem likely candidates as adjuncts in a multimodal approach to postoperative pain control. They produce small and most often clinically minor changes in respiratory function and importantly, they do not usually exacerbate opioid-induced respiratory depression. Caution is appropriate as occasionally, significant respiratory depression or hypotension may still result.
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Time (min) Figure 13.17 Average ventilation at a PETCO2 ¼ 55 mm Hg as determined by the hypercapnic ventilatory response. For clarity, SEM is shown for the placebo and 0.5 and 1.0 mg kg1 groups. See Figure 13.1 for explanation of the measurement time points. P 5 .05 different from the baseline (time ¼ 0) measurement within a dose þ group P 5 .05 different from the placebo measurement at the same time (Data from Ref. 294).
D. N-Methyl-D-Aspartate (NMDA) Receptor Antagonists Dextromethorphan
Activation of N-methyl-D-aspartate (NMDA) receptors plays a role in central sensitization after peripheral tissue injury, wind-up phenomenon [311], and the development of opioid tolerance [312]. Pre-emptive treatment with dextromethorphan, compared with the same treatment at the end of surgery, has been shown to reduce opioid requirement and reduce the incidence of postoperative hypoxemia and nausea [312]. Ketamine
Ketamine is a noncompetitive N-methyl-D-aspartate (NMDA) receptor antagonist [313] and also mediates some analgesia via opioid receptors [314]. Ketamine also may have sodium and potassium channel effects similar to local anesthetics in nerve membranes [315]. Ketamine can produce significant analgesia and is thought to stimulate ventilation, and thus, it can attenuate the hypoventilation induced by other drugs [316]. However, this potentially beneficial effect may not be predictable or consistent. Ketamine alone usually has minimal effects on central respiratory drive measured by CO2 responsiveness [317]. Occasionally, hypoventilation
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or apnea can follow bolus doses of ketamine [318]. Thus ketamine may have the potential to depress respiration, most likely mediated by mu receptors, especially when combined with other drugs. Ketamine is also a bronchodilator and can improve pulmonary compliance and break bronchospastic episodes resistant to conventional therapy [319]. Ketamine may help maintain airway function but it can also increase upper airway reactivity. Even small doses of ketamine, as part of a multimodal approach to postoperative analgesia, can improve dynamic analgesia [320]. Intravenous ketamine (0.5 mg kg1 bolus plus 0.25 mg kg1 h1) has been reported to reduce wound hyperalgesia for up to six months after surgery for rectal adenocarcinoma [321]. Intravenous ketamine was superior to epidural ketamine in this regard. Others report that adding ketamine to morphine for PCA analgesia after abdominal surgery did not improve analgesia but did lead to vivid dreams and worse performance on tests of cognitive function [322]. Epidural ketamine has been reported to improve analgesia after upper abdominal surgery when compared with epidural morphine alone [323]. Intrathecal ketamine added to bupivacaine did not prolong postoperative analgesia after intracavitary brachytherapy when compared with bupivacaine alone, but did increase side effects (nausea, vomiting, dizziness, sedation, and strange feelings) [324]. Others have also found intrathecal ketamine either not to be consistently effective or to have too significant a side effect profile. The efficacy of intrathecal ketamine remains uncertain. Although respiratory depression has not been raised as a concern with intrathecal ketamine, issues over potential neurotoxicity, as well as considerable psychomimetic and cardiovascular side effects, currently limit its use via this route. In summary, small doses of ketamine may very well play a role in balanced or multi-modal analgesia. It is possible that superior NMDA antagonists will be available in the future. For now, although potentially beneficial as an analgesic adjunct, ketamine can produce unwanted side effects, including increased secretions, cardiovascular stimulation, and psychomimetic effects. These can range from vivid dreams to overt psychotic hallucinations that can be frightening to individuals experiencing them as well as to family and friends. E.
Nonsteroidal Anti-inflammatory Drugs (NSAIDs)
New potent NSAIDs can produce significant analgesia shortly after intravenous injection. They are also associated with potentially significant adverse effects, including renal dysfunction or failure, platelet function inhibition, and gastrointestinal bleeding. NSAIDs can be administered rectally or intravenously. The appropriate use of NSAIDs for postoperative
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pain relief is not usually associated with increased bleeding or renal dysfunction. NSAIDs are not reported to produce respiratory depression and can be combined with other analgesic therapies to contribute to multimodal analgesia. Frequently, opioid requirements (and presumably associated side effects) can be reduced approximately 20–30% [325] by the co-administration of NSAIDs. NSAIDs, when combined with opioids to control postoperative pain, may not always reduce opioid-induced side effects, including respiratory depression [326]. No studies allow identification and isolation of outcome benefits attributable to NSAIDs [21]. It is not clear if adding systemic NSAIDs to pain control using thoracic epidural analgesia with local anesthetic and opioid is useful [212]. F. Neostigmine
Epidural neostigmine has been reported to prolong the analgesia provided by epidural bupivacaine after abdominal hysterectomy [327]. Both clonidine and neostigmine also have been reported to produce analgesia after intraarticular injection in patients undergoing knee surgery [305]. Intrathecal neostigmine, a cholinesterase inhibitor, produces analgesia in humans [328]. Neostigmine, especially when combined with other drugs, may promote hemodynamic stability because it stimulates sympathetic preganglionic neurons [329]. The role of intrathecal neostigmine, as well as the potential for respiratory adverse effects or complications when it is used, has not been studied. VIII.
Pre-Emptive Analgesia
Reviews of pre-emptive analgesia are available in the literature [330]. The validity and significance of pre-emptive analgesia continues to be debated. It is generally agreed that if pre-emptive analgesia is truly to be advantageous, then dynamic analgesia should be achieved and consistently maintained throughout the perioperative period [69]. Few studies have examined this aspect of pain control; outcomes and results have been conflicting [18,331]. To date, a few studies have demonstrated improved short- and intermediate-term analgesia with pre-emptive analgesia [18,331]. IX.
Perioperative Analgesia and Pulmonary Outcome
Studies by Yeager et al. [20] and Tuman et al. [63] highlighted how different approaches to anesthesia and analgesia might modify and improve patient outcome. In this chapter, several key points have been established concerning the roles that surgery and pain play in contributing to postoperative changes in respiratory function and how effective pain relief
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may achieve improvements in postoperative pulmonary outcome. Among the most important of these are the following: 1.
2.
3.
4.
5.
6.
Surgical pain contributes to, but is not the only cause of, respiratory and pulmonary dysfunction after surgery. Other inhibitory reflexes play important roles. It should not be expected that good pain control in and of itself (as can be achieved with opioids) would be sufficient to ameliorate respiratory dysfunction after surgery. Respiratory and pulmonary dysfunction is most impaired after upper abdominal or thoracic surgery, and it is in these surgical candidates, as well as in high-risk patients, where the most gain is likely to be achieved with more complex analgesic strategies in an attempt to reduce postoperative morbidity. Optimal analgesia should be multimodal, pre-emptive, continuous, consistent, and dynamic. This will allow reduction of adverse effects; it will provide maximal inhibition and/or suppression of both surgical pain and associated deleterious reflexes and responses, and the best chance for real outcome improvement. Traditional surrogate measures of pulmonary function (lung function studies, e.g., FEV1) do not allow benefit prediction (see below). It is perhaps time to accept that current use of these surrogate measures is inadequate and that other tools (e.g., measurements of diaphragmatic function) should be studied and developed. Epidural analgesia is technically challenging. Practical problems with epidural analgesia can override potential benefits and impede successful pain control [7]. In addition, combining opioids and local anesthetics, especially in higher concentrations or greater doses, can cause morbidity [332].
Epidural analgesia with a combination of local anesthetic and opioid is widely perceived as the preferred technique for postoperative pain control [333]. A significant number of reviews and meta-analyses of over 100 studies identify the benefits of regional anesthesia or analgesia (see below) [11,18,19,21,334]. Nevertheless, all too frequently, epidural analgesia is not administered. Factors cited as contributing to this underutilization of a supposedly optimal technique include concerns about safety, time, challenges and difficulties in effectively operating a pain service, and doubts about the real benefits of epidural analgesia. McLeod et al. [7] have documented the problems with delivering effective epidural analgesia. Of 640 patients receiving epidural analgesia, 20% did not have satisfactory pain control. Almost one-third of these patients had a problem with the
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epidural, and technical problems were experienced in 13% of patients. Rigg et al. [6] reported that fully 50% of more than 900 patients randomized to receive epidural analgesia for 72 h postoperatively had some protocol violation that most commonly was related to an epidural catheter failure. Approximately 10% of patients experienced problems with catheter insertion or immediate efficacy. The difficulties involved in establishing whether regional analgesia improves patient outcomes are significant and have recently been reviewed [18,42]. If regional analgesia is to become a standard of care, resources must be garnered to support it. This will require the extension of proven outcome benefits beyond traditional clinical end-points (e.g., atelectasis, pneumonia) to functional health status, patient satisfaction, and economic impact. Large sample sizes are required to demonstrate improvements in uncommon outcomes. Randomized controlled trials, considered to be among the best study designs, cannot always achieve the required sample size for certain end-points. Meta-analyses, recognized to have some limitations, have become a popular tool for combining numerous independently performed investigations [18]. Ballantyne et al. [11] reported a meta-analysis of randomized controlled trials evaluating the effects of postoperative analgesic therapies on pulmonary outcome. Their efforts stemmed from those of the Agency for Health Care Policy and Research to establish which medical practices deserve strong endorsement. The meta-analysis originally retrieved 121 studies from MEDLINE (none of the other 11 databases searched contributed additional relevant studies), of which 65 studies, dating from 1966 to 1995, were used. Concerning epidural opioids versus systemic opioids (24 studies): pain relief was better with epidural opioids in nine of 24 studies; pooled data revealed atelectasis to be significantly less with epidural opioids even though only three individual studies had this finding. There was a trend toward a reduction in pulmonary complications with epidural versus systemic opioids and surrogate measures of pulmonary function (e.g., FEV1) did not reach significance between the two approaches when data was pooled. Concerning epidural local anesthetics versus systemic opioids (11 studies): the incidence of pulmonary infections and other complications was reduced with epidural local anesthetic versus systemic opioids; again, surrogate measures of pulmonary function were not different between the two treatment groups. Concerning epidural local anesthetic with opioids versus systemic opioids (seven studies): analgesia was superior in all seven trials, and oxygenation was better with epidural local anesthetic plus opioid analgesia, though few studies controlled for supplemental oxygen administration, rendering interpretation of this finding difficult. No conclusions concerning the predictive value of surrogate measures of pulmonary function or the incidence of pulmonary
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complications could be made due either to design of the individual trials or to inadequate number of trials. Concerning thoracic versus lumbar epidural opioids (eight studies): there were no significant differences between the two treatment approaches. Kehlet and Holte recently reviewed the effect of postoperative analgesia on surgical outcome [21], reporting that continuous epidural analgesia with local anesthetic reduced pulmonary complications from 16.7% to 10.4% in patients after major abdominal or vascular surgery. When only considering studies of thoracic epidural analgesia, the reduction in pulmonary complications (16.7–10.9%) did not reach statistical significance. In thoracic surgery studies, where thoracic epidurals most commonly included opioids, a significant reduction in pulmonary complications was achieved (31.1–14.6%). These results were largely derived from two of the 12 studies considered in this sub-analysis. Kehlet and Holte [21] concluded that continuous epidural local anesthetic or local anestheticopioid analgesia has only been demonstrated to reduce postoperative pulmonary morbidity after major abdominal surgery. They note that more studies are required to validate results, suggesting that epidural opioids or opioid-local anesthetic mixtures can produce significant pulmonary morbidity reduction after surgery. Since the above-cited meta-analyses, additional studies continue to appear in the literature addressing the matter of anesthetic and/or analgesic technique and patient outcome. Park et al. [334] reported a randomized trial of 1,021 patients undergoing abdominal surgery who received either general anesthesia with parenteral opioid analgesia postoperatively versus combined epidural-general anesthesia with epidural morphine for postoperative analgesia. They found a significant difference in death or major complications only in the subset of patients who had abdominal aortic surgery. There was a trend for a reduction in the number of patients who had respiratory failure in the entire study group (71 versus 51). Respiratory failure was significantly reduced in abdominal aortic surgery patients receiving epidural anesthesia/analgesia (n ¼ 26) versus those who did not (n ¼ 52). The incidence of pneumonia was less but not statistically significant (8 versus 19, p ¼ 0.06). Respiratory depression did seem more common in the epidural group (14 versus 6) but the difference was not statistically significant. Intraoperative use of epidural anesthesia and postoperative analgesia also was reported to result in improved oxygenation, more stable cardiac output, earlier extubation, better pain control and shorter ICU stay in patients undergoing lung resection [335]. Rigg et al. [6] reported results of the MASTER trial (Multicentre Australian Study of Epidural Anaesthesia) where 915 patients undergoing major abdominal surgery were randomized to receive either general anesthesia with routine postoperative analgesia (e.g., PCA) or epidural anesthesia and epidural analgesia (local anesthetic plus opioid) for 72 h postoperatively. Twenty-five centers participated but
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it still took five years for an adequate number of subjects to be enrolled according to the power analysis. Pain scores were better in the epidural group but of all the morbidity categories, only differences in respiratory failure reached statistical significance. Respiratory failure was defined as the need for prolonged ventilation, the need for reintubation, or PaO2 5 51 mm Hg or PaCO2 4 49 mm Hg with patients breathing room air. Unfortunately, inconsistent findings continue to be documented. Norris et al. [336] studied 168 abdominal aorta surgery patients randomized to one of four groups where operative anesthesia was either general or epidural plus ‘light’ general and postoperative analgesia was either IV or epidural PCA for at least 72 h. Except for a small (3–6 h) difference in the time to tracheal extubation, the authors found no difference in outcomes which included length of stay, death, cardiovascular and respiratory complications, and other common clinical endpoints.
X.
Cardiac Surgery
Although surgical pain is less after sternotomy for cardiac surgery when compared with thoracotomy, pulmonary morbidity remains significant after cardiac surgery. Typically, a restrictive defect develops after surgery and can persist for days or more. Reductions in lung volume and other spirometric values can exceed 35% [337]. Decreases in FVC, FEV1 and PEFR are common, ranging from 30% to 60%, and can persist for more than five days, the current average length of stay for many cardiac operations [338]. Predictable decreases in oxygenation, stemming from ventilation–perfusion mismatch and shunt also occur. Multifactorial in origin, factors such as muscle weakness, pain, thoraco-abdominal wall motion abnormalities, interstitial edema, diaphragmatic paresis, pleural effusions, bleeding within the chest, and effects of pleurotomy all play roles. How the internal thoracic (mammary) artery is surgically prepared as a graft also affects pulmonary function. Dissection of all the surrounding tissue with opening of the pleura, when compared with preparing a venoarterial pedicle leaving the pleura intact, can result in lower lung function after surgery. Common respiratory complications after cardiac surgery include persistent hypoxemia, atelectasis, pneumonia, and respiratory failure with ventilator dependency. Numerous studies have attempted to optimize pain relief after cardiac surgery. Initial investigations in the 1980s and 1990s evaluated intrathecal morphine [339–341]. Relatively low doses of intrathecal morphine (e.g., 5 mg kg1) appear to minimize side effects and produce effective postoperative analgesia after cardiac surgery. However, little consistent benefit has been proven to be associated with intrathecal morphine analgesia after cardiac surgery. Intermediate outcomes, such as
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the time to tracheal extubation, can actually be prolonged. No studies have demonstrated intrathecal morphine analgesia to be associated with a decrease in pulmonary complications. More recently, thoracic epidural analgesia for patients undergoing cardiac surgery has received attention. Thoracic epidural analgesia with local anesthetics and opioids can provide intense analgesia, block sympathetic reflexes that are potentially deleterious to myocardial oxygen supply and demand ratios, and improve coronary circulation [215,342]. Laboratory and human studies suggest that myocardial stunning, ischemia, and infarct size may potentially be reduced with thoracic epidural analgesia. Other purported benefits of thoracic epidural analgesia in patients undergoing cardiac surgery include excellent analgesia with near elimination of the need for opioids and their associated side effects, improved coronary blood flow, early tracheal extubation, and reductions in renal failure, arrhythmias, and confusion [343]. Several reports claim improvement in pulmonary outcome with minimal to no respiratory complications related to analgesia [344–346]. Concerns over epidural instrumentation and catheterization in patients who require anticoagulation for cardiopulmonary bypass, along with the need to carefully screen patients and to dedicate the time and technical expertise required to insert and properly maintain analgesia with a thoracic epidural, severely limit enthusiasm for this mode of analgesia in cardiac surgical patients. Additional questions, such as whether discontinuing platelet inhibiting agents (including aspirin) for days to weeks prior to surgery is feasible or safe, remain unanswered. Nevertheless, evidence is accumulating that thoracic epidural analgesia can be both safe and effective in cardiac surgical patients. To date over 6,000 cardiac surgical patients have been cumulatively reported to have received thoracic epidural analgesia for cardiac surgery without a single spinal hematoma being reported [346]. Nevertheless, without firm evidence that thoracic epidural analgesia produces cost-effective benefit that cannot be achieved by other means, the widespread application of this therapy in cardiac surgical patients is not likely to become popular. Fewer investigations have sought to determine if there are reductions in pulmonary complications associated with the use of thoracic epidural analgesia in cardiac surgical patients [232,347]. These studies were not sufficiently designed to permit acceptance of their purported claims that epidural analgesia in cardiac surgical patients leads to improvements in intermediate outcomes such as earlier tracheal extubation, better oxygenation, and improved lung volume and flow variables. Others also have reported similar results in patients undergoing cardiac surgery; namely, better pain relief and earlier extubation but no difference in the length of hospital stay [348]. Additional large randomized trials to examine both the benefit and risk of thoracic epidural analgesia after cardiac surgery are required.
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Summary
Major (e.g., abdominal, thoracic) surgery produces significant pain and severely impairs respiratory function. Dynamic, multi-modal analgesia that includes regional analgesia with low doses of local anesthetics and opioids can provide optimal pain relief and permit aggressive rehabilitation and physical therapy. Properly administered analgesic therapies carry a low incidence of clinically significant respiratory depression that can be effectively monitored, detected and treated, preventing harm. Combined with early ambulation and nutrition, such approaches offer the best chances for reducing pulmonary morbidity as well as other complications after surgery. However, it is not simple to successfully achieve analgesia with approaches that include epidural techniques (Table 13.4). Only a dedicated, well-organized service, such as an acute pain service, should be expected to be expert and vigilant to the degree required to optimize efficacy and safety of postoperative analgesia. Multi-modal analgesia [67] should include or consider each of the following: 1. 2. 3.
4.
5. 6. 7.
8. 9.
10.
Patient education concerning postoperative goals and care is important. Use of pre-emptive analgesia strategies (e.g., NMDA antagonists) can reduce certain aspects of the pain response. Regional analgesia is an integral approach to postoperative pain control, which will produce dynamic analgesia and is best placed at or near the site of injury. Regional analgesia should employ low concentrations of local anesthetic and low doses of opioid in order to produce optimal analgesia with minimal side effects. Other adjuncts that are being explored as regional analgesics and modulators of responses to noxious stimulation or trauma should be considered. Use of systemic NSAIDs and other analgesics with fewer adverse effects than opioids can be opioid sparing. Systemic or parenteral opioids should be prescribed as needed for short-term analgesia. It should be recognized that the risk of respiratory depression with opioid-based analgesic techniques is approximately 1% regardless of the route of drug administration. Supplemental oxygen is inexpensive, very safe, and effectively reduces hypoxemia. The use of prophylactic low-dose naloxone infusions to reduce opioid-induced side effects, no matter what their route of administration, should be considered. Dynamic analgesia will allow appropriately aggressive physical therapy, ambulation and expeditious recovery.
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Table 13.4 Patient
Regimen
Staff
Monitoring
Audit
Effective and Safe Management of Ward-based Epidural Analgesia Informed consent Careful selection based on a risk/benefit analysis and absence of contraindications Sterile technique Standard, pharmacy prepared or commercially produced low-dose LA-opioid Infusion Bupivacaine 8–12 mg h1 plus fentanyl 2–4 mg ml1 or morphine/diamorphine 50 mg ml1 Standard infusion pump (different from IV PCA pumps) with PCEA facility Identifiable administration sets without injection ports Bacterial filter Transparent dressing Training program/protocols/acute pain handbook with particular attention to: Recognition and management of complications including hypotension, respiratory depression, inadequate analgesia and motor blockade Concurrent thromboprophylaxis and anticoagulant therapy Access to members of the acute pain team 24 h daily Regular monitoring of dynamic pain scores, cardiorespiratory parameters, sedation scores, dermatomal level and motor blockade Daily inspection of the epidural site Twice daily review by the APT Audit and feedback to anesthetists, surgeons and nurses Critical incident reporting
Other obstacles to producing optimal analgesia and reducing postoperative morbidity include: (1) the need for additional evidence of benefit; (2) evidence of which patient populations will benefit; (3) challenges in efficiently and effectively producing regional analgesia, and (4) finding the resources to allow the pursuit of the above. References 1.
Marks, R.M. and Sachar, E.J., Undertreatment of medical inpatients with narcotic analgesics, Ann. Intern. Med. 78, 173–181, 1973. 2. Cohen, F.L., Postsurgical pain relief: Patients’ status and nurses’ medication choices, Pain 9, 265–274, 1980. 3. Ward, S.E. and Gordon, D., Application of the American Pain Society quality assurance standards, Pain 56, 299–306, 1994.
Pain Management and Regional Anesthesia 4. 5. 6.
7.
8. 9.
10.
11.
12. 13. 14. 15.
16. 17.
18. 19.
20.
21.
491
Svensson, I., Sjostrom, B. and Haljamae, H., Assessment of pain experiences after elective surgery, J. Pain Symptom Manag. 20, 193–201, 2000. Schafheutle, E.I., Cantrill, J.A. and Noyce, P.R., Why is pain management suboptimal on surgical wards? J. Adv. Nurs. 33, 728–737, 2001. Rigg, J.R., Jamrozik, K., Myles, P.S., Silbert, B.S., Peyton, P.J., Parsons, R.W. and Collins, K.S., Epidural anaesthesia and analgesia and outcome of major surgery: A randomised trial, Lancet 359, 1276–1282, 2002. McLeod, G., Davies, H., Munnoch, N., Bannister, J. and MacRae, W., Postoperative pain relief using thoracic epidural analgesia: Outstanding success and disappointing failures, Anaesthesia 56, 75–81, 2001. Wheatley, R.G., Schug, S.A. and Watson, D., Safety and efficacy of postoperative epidural analgesia, Br. J. Anaesth. 87, 47–61, 2001. Kavanagh, B.P., Katz, J. and Sandler, A.N., Pain control after thoracic surgery. A review of current techniques, Anesthesiology 81, 737–759, 1994. Eisenberg, E., Pultorak, Y., Pud, D. and Bar-El, Y., Prevalence and characteristics of post coronary artery bypass graft surgery pain (PCP), Pain 92, 11–17, 2001. Ballantyne, J.C., Carr, D.B., deFerranti, S., Suarez, T., Lau, J., Chalmers, T.C., Angelillo, I.F. and Mosteller, F., The comparative effects of postoperative analgesic therapies on pulmonary outcome: Cumulative meta-analyses of randomized, controlled trials, Anesth. Analg. 86, 598–612, 1998. Warner, D.O., Preventing postoperative pulmonary complications: The role of the anesthesiologist, Anesthesiology 92, 1467–1472, 2000. Craig, D.B., Postoperative recovery of pulmonary function, Anesth. Analg. 60, 46–52, 1981. Marshall, B.E. and Wyche, M.Q., Jr., Hypoxemia during and after anesthesia, Anesthesiology 37, 178–209, 1972. Pavlin, E.G., Holle, R.H. and Schoene, R.B., Recovery of airway protection compared with ventilation in humans after paralysis with curare, Anesthesiology 70, 381–385, 1989. Nunn, J.F., Effects of anaesthesia on respiration, Br. J. Anaesth. 65, 54–62, 1990. Nishino, T., Hiraga, K., Fujisato, M., Mizuguchi, T. and Honda, Y., Breathing patterns during postoperative analgesia in patients after lower abdominal operations, Anesthesiology 69, 967–972, 1988. Wu, C.L. and Fleisher, L.A., Outcomes research in regional anesthesia and analgesia, Anesth. Analg. 91, 1232–1242, 2000. Rodgers, A., Walker, N., Schug, S., McKee, A., Kehlet, H., van Zundert, A., Sage, D., Futter, M., Saville, G., Clark, T. and MacMahon, S., Reduction of postoperative mortality and morbidity with epidural or spinal anaesthesia: Results from overview of randomised trials, BMJ 321, 1493, 2000. Yeager, M.P., Glass, D.D., Neff, R.K. and Brinck-Johnsen, T., Epidural anesthesia and analgesia in high-risk surgical patients, Anesthesiology 66, 729–736, 1987. Kehlet, H. and Holte, K., Effect of postoperative analgesia on surgical outcome, Br. J. Anaesth. 87, 62–72, 2001.
492 22. 23. 24.
25. 26.
27.
28.
29.
30.
31.
32. 33.
34. 35.
36.
37.
Bailey and Thakur Weissman, C., Pulmonary function after cardiac and thoracic surgery, Anesth. Analg. 88, 1272–1279, 1999. Sandler, A.N., Post-thoracotomy analgesia and perioperative outcome, Minerva Anestesiol. 65, 267–274, 1999. Catley, D.M., Thornton, C., Jordan, C., Lehane, J.R., Royston, D. and Jones, J.G., Pronounced, episodic oxygen desaturation in the postoperative period: Its association with ventilatory pattern and analgesic regimen, Anesthesiology 63, 20–28, 1985. Liu, S., Carpenter, R.L. and Neal, J.M., Epidural anesthesia and analgesia. Their role in postoperative outcome, Anesthesiology 82, 1474–1506, 1995. Fratacci, M.D., Kimball, W.R., Wain, J.C., Kacmarek, R.M., Polaner, D.M. and Zapol, W.M., Diaphragmatic shortening after thoracic surgery in humans. Effects of mechanical ventilation and thoracic epidural anesthesia, Anesthesiology 79, 654–665, 1993. Dureuil, B., Cantineau, J.P. and Desmonts, J.M., Effects of upper or lower abdominal surgery on diaphragmatic function, Br. J. Anaesth. 59, 1230–1235, 1987. Ali, J., Weisel, R.D., Layug, A.B., Kripke, B.J. and Hechtman, H.B., Consequences of postoperative alterations in respiratory mechanics, Am. J. Surg. 128, 376–382, 1974. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G., Reber, A. and Hedenstierna, G., Prevention of atelectasis during general anaesthesia, Lancet 345, 1387–1391, 1995. Edmark, L., Kostova-Aherdan, K., Enlund, M. and Hedenstierna, G., Optimal oxygen concentration during induction of general anesthesia, Anesthesiology 98, 28–33, 2003. Mangano, D.T. and Goldman, L., Preoperative assessment of patients with known or suspected coronary disease, N. Engl. J. Med. 333, 1750–1756, 1995. Garibaldi, R.A., Britt, M.R., Coleman, M.L., Reading, J.C. and Pace, N.L., Risk factors for postoperative pneumonia, Am. J. Med. 70, 677–680, 1981. Rosenberg, J., Dirkes, W.E. and Kehlet, H., Episodic arterial oxygen desaturation and heart rate variations following major abdominal surgery, Br. J. Anaesth. 63, 651–654, 1989. Peters, R.M., Wellons, H.A., Jr. and Htwe, T.M., Total compliance and work of breathing after thoracotomy, J. Thorac. Cardiovasc. Surg. 57, 348–355, 1969. Nomori, H., Horio, H., Fuyuno, G. and Kobayashi, R., Non-serratus-sparing antero-axillary thoracotomy with disconnection of anterior rib cartilage. Improvement in postoperative pulmonary function and pain in comparison to posterolateral thoracotomy, Chest 111, 572–576, 1997. Pedersen, T., Eliasen, K. and Henriksen, E., A prospective study of risk factors and cardiopulmonary complications associated with anaesthesia and surgery: Risk indicators of cardiopulmonary morbidity, Acta Anaesthesiol. Scand. 34, 144–155, 1990. Joris, J., Kaba, A. and Lamy, M., Postoperative spirometry after laparoscopy for lower abdominal or upper abdominal surgical procedures, Br. J. Anaesth. 79, 422–426, 1997.
Pain Management and Regional Anesthesia 38.
39. 40. 41.
42. 43. 44.
45.
46.
47.
48. 49.
50.
51.
52. 53.
493
Furrer, M., Rechsteiner, R., Eigenmann, V., Signer, C., Althaus, U. and Ris, H.B., Thoracotomy and thoracoscopy: Postoperative pulmonary function, pain and chest wall complaints, Eur. J. Cardiothorac. Surg. 12, 82–87, 1997. Weissman, C., The metabolic response to stress: An overview and update, Anesthesiology 73, 308–327, 1990. Kehlet, H., Surgical stress: The role of pain and analgesia, Br. J. Anaesth. 63, 189–195, 1989. Modig, J., Respiration and circulation after total hip replacement surgery. A comparison between parenteral analgesics and continuous lumbar epidural block, Acta Anaesthesiol. Scand. 20, 225–236, 1976. Breivik, H., Postoperative pain management: Why is it difficult to show that it improves outcome? Eur. J. Anaesthesiol. 15, 748–751, 1998. Kehlet, H., The surgical stress response: Should it be prevented? Can. J. Surg. 34, 565–567, 1991. Yeager, M.P., Yu, C.T., Campbell, A.S., Moschella, M. and Guyre, P.M., Effect of morphine and beta-endorphin on human Fc receptor-dependent and natural killer cell functions, Clin. Immunol. Immunopathol. 62, 336–343, 1992. Ryhanen, P., Jouppila, R., Lanning, M., Jouppila, P., Hollmen, A. and Kouvalainen, K., Natural killer cell activity after elective cesarean section under general and epidural anesthesia in healthy parturients and their newborns, Gynecol. Obstet. Investig. 19, 139–142, 1985. Anand, K.J. and Hickey, P.R., Halothane–morphine compared with high-dose sufentanil for anesthesia and postoperative analgesia in neonatal cardiac surgery (see comments), N. Engl. J. Med. 326, 1–9, 1992. Brodner, G., Van Aken, H., Hertle, L., Fobker, M., Von Eckardstein, A., Goeters, C., Buerkle, H., Harks, A. and Kehlet, H., Multimodal perioperative management—combining thoracic epidural analgesia, forced mobilization, and oral nutrition—reduces hormonal and metabolic stress and improves convalescence after major urologic surgery, Anesth. Analg. 92, 1594–1600, 2001. Wahba, W.M., Don, H.F. and Craig, D.B., Post-operative epidural analgesia: Effects on lung volumes, Can. Anaesth. Soc. J. 22, 519–527, 1975. Cuschieri, R.J., Morran, C.G., Howie, J.C. and McArdle, C.S., Postoperative pain and pulmonary complications: Comparison of three analgesic regimens, Br. J. Surg. 72, 495–498, 1985. Clergue, F., Montembault, C., Despierres, O., Ghesquiere, F., Harari, A. and Viars, P., Respiratory effects of intrathecal morphine after upper abdominal surgery, Anesthesiology 61, 677–685, 1984. Manikian, B., Cantineau, J.P., Bertrand, M., Kieffer, E., Sartene, R. and Viars, P., Improvement of diaphragmatic function by a thoracic extradural block after upper abdominal surgery, Anesthesiology 68, 379–386, 1988. Mangano, D.T., Perioperative cardiac morbidity, Anesthesiology 72, 153–184, 1990. Hollenberg, M., Mangano, D.T., Browner, W.S., London, M.J., Tubau, J.F. and Tateo, I.M., Predictors of postoperative myocardial ischemia in patients undergoing noncardiac surgery. The Study of Perioperative Ischemia Research Group, JAMA 268, 205–209, 1992.
494 54.
55. 56.
57.
58.
59.
60. 61.
62.
63.
64. 65.
66.
67. 68.
69.
Bailey and Thakur Reiz, S., Haggmark, S., Rydvall, A. and Ostman, M., Beta-blockers and thoracic epidural analgesia. Cardioprotective and synergistic effects, Acta Anaesthesiol. Scand. Suppl 76, 54–61, 1982. Landesberg, G., The pathophysiology of perioperative myocardial infarction: Facts and perspectives, J. Cardiothorac. Vasc. Anesth. 17, 90–100, 2003. Beattie, W.S., Buckley, D.N. and Forrest, J.B., Epidural morphine reduces the risk of postoperative myocardial ischaemia in patients with cardiac risk factors, Can. J. Anaesth. 40, 532–541, 1993. Breslow, M.J., Jordan, D.A., Christopherson, R., Rosenfeld, B., Miller, C.F., Hanley, D.F., Beattie, C., Traystman, R.J. and Rogers, M.C., Epidural morphine decreases postoperative hypertension by attenuating sympathetic nervous system hyperactivity, JAMA 261, 3577–3581, 1989. Beattie, W.S., Badner, N.H. and Choi, P., Epidural analgesia reduces postoperative myocardial infarction: A meta-analysis, Anesth. Analg. 93, 853– 858, 2001. Parker, S.D., Breslow, M.J., Frank, S.M., Rosenfeld, B.A., Norris, E.J., Christopherson, R., Rock, P., Gottlieb, S.O., Raff, H., Perler, B.A., et al. Catecholamine and cortisol responses to lower extremity revascularization: Correlation with outcome variables. Perioperative Ischemia Randomized Anesthesia Trial Study Group, Crit. Care Med. 23, 1954–1961, 1995. Cousins, M.J., John J. Bonica distinguished lecture. Acute pain and the injury response: Immediate and prolonged effects, Reg. Anesth. 14, 162–179, 1989. Perhoniemi, V. and Linko, K., Effect of spinal versus epidural anaesthesia with 0.5% bupivacaine on lower limb blood flow, Acta Anaesthesiol. Scand. 31, 117– 121, 1987. Henny, C.P., Odoom, J.A., ten Cate, H., ten Cate, J.W., Oosterhoff, R.J., Dabhoiwala, N.F. and Sih, I.L., Effects of extradural bupivacaine on the haemostatic system, Br. J. Anaesth. 58, 301–305, 1986. Tuman, K.J., McCarthy, R.J., March, R.J., DeLaria, G.A., Patel, R.V. and Ivankovich, A.D., Effects of epidural anesthesia and analgesia on coagulation and outcome after major vascular surgery, Anesth. Analg. 73, 696–704, 1991. Steinbrook, R.A., Epidural anesthesia and gastrointestinal motility, Anesth. Analg. 86, 837–844, 1998. Ahn, H., Bronge, A., Johansson, K., Ygge, H. and Lindhagen, J., Effect of continuous postoperative epidural analgesia on intestinal motility, Br. J. Surg. 75, 1176–1178, 1988. Scheinin, B., Asantila, R. and Orko, R., The effect of bupivacaine and morphine on pain and bowel function after colonic surgery, Acta Anaesthesiol. Scand. 31, 161–164, 1987. Kehlet, H., Multimodal approach to control postoperative pathophysiology and rehabilitation, Br. J. Anaesth. 78, 606–617, 1997. Rady, M.Y., Ryan, T. and Starr, N.J., Early onset of acute pulmonary dysfunction after cardiovascular surgery: Risk factors and clinical outcome, Crit. Care Med. 25, 1831–1839, 1997. Woolf, C.J. and Chong, M.S., Preemptive analgesia—treating postoperative pain by preventing the establishment of central sensitization, Anesth. Analg. 77, 362–379, 1993.
Pain Management and Regional Anesthesia 70. 71.
72. 73.
74.
75.
76.
77. 78. 79. 80.
81.
82. 83.
84. 85.
86.
87.
495
Kalso, E., Perttunen, K. and Kaasinen, S., Pain after thoracic surgery, Acta Anaesthesiol. Scand. 36, 96–100, 1992. Obata, H., Saito, S., Fujita, N., Fuse, Y., Ishizaki, K. and Goto, F., Epidural block with mepivacaine before surgery reduces long-term post-thoracotomy pain, Can. J. Anaesth. 46, 1127–1132, 1999. Millan, M.J., The induction of pain: An integrative review, Prog. Neurobiol., 57, 1–164, 1999. Waldrop, T.G., Millhorn, D.E., Eldridge, F.L. and Klingler, L.E., Respiratory responses to noxious and nonnoxious heating of skin in cats, J. Appl. Physiol. 57, 1738–1741, 1984. Duranti, R., Pantaleo, T., Bellini, F., Bongianni, F. and Scano, G., Respiratory responses induced by the activation of somatic nociceptive afferents in humans, J. Appl. Physiol. 71, 2440–2448, 1991. Sarton, E., Dahan, A., Teppema, L., Berkenbosch, A., van den Elsen, M. and van Kleef, J., Influence of acute pain induced by activation of cutaneous nociceptors on ventilatory control, Anesthesiology 87, 289–296, 1997. Borgbjerg, F.M., Nielsen, K. and Franks, J., Experimental pain stimulates respiration and attenuates morphine-induced respiratory depression: A controlled study in human volunteers, Pain 64, 123–128, 1996. Bourke, D.L., Respiratory effects of regional anesthesia during acute pain, Reg. Anesth. 18, 361–365, 1993. Lambert, D.G., Recent advances in opioid pharmacology, Br. J. Anaesth. 81, 1–2, 1998. Pert, C.B. and Snyder, S.H., Opiate receptor: Demonstration in nervous tissue, Science 179, 1011–1014, 1973. Terenius, L., Characteristics of the ‘receptor’ for narcotic analgesics in synaptic plasma membrane fraction from rat brain, Acta Pharmacol. Toxicol. (Copenh.) 33, 377–384, 1973. Simon, E.J., Hiller, J.M. and Edelman, I., Stereospecific binding of the potent narcotic analgesic (3H) Etorphine to rat-brain homogenate, Proc. Natl. Acad. Sci. USA 70, 1947–1949, 1973. Bonham, A.C., Neurotransmitters in the CNS control of breathing, Respir. Physiol. 101, 219–230, 1995. Shook, J.E., Watkins, W.D. and Camporesi, E.M., Differential roles of opioid receptors in respiration, respiratory disease, and opiate-induced respiratory depression, Am. Rev. Respir. Dis. 142, 895–909, 1990. Santiago, T.V. and Edelman, N.H., Opioids and breathing, J. Appl. Physiol. 59, 1675–1685, 1985. Sora, I., Takahashi, N., Funada, M., Ujike, H., Revay, R.S., Donovan, D.M., Miner, L.L. and Uhl, G.R., Opiate receptor knockout mice define mu receptor roles in endogenous nociceptive responses and morphine-induced analgesia, Proc. Natl. Acad. Sci. USA 94, 1544–1549, 1997. Ling, G.S. and Pasternak, G.W., Spinal and supraspinal opioid analgesia in the mouse: The role of subpopulations of opioid binding sites, Brain Res. 271, 152–156, 1983. Dahan, A., Sarton, E., Teppema, L., Olievier, C., Nieuwenhuijs, D., Matthes, H.W. and Kieffer, B.L., Anesthetic potency and influence of morphine
496
88. 89. 90. 91.
92.
93.
94. 95.
96.
97.
98.
99.
100.
101.
102.
103.
Bailey and Thakur and sevoflurane on respiration in mu-opioid receptor knockout mice, Anesthesiology 94, 824–832, 2001. Pasternak, G.W. and Snyder, S.H., Identification of novel high affinity opiate receptor binding in rat brain, Nature 253, 563–565, 1975. Pick, C.G., Paul, D. and Pasternak, G.W., Nalbuphine, a mixed kappa 1 and kappa 3 analgesic in mice, J. Pharmacol. Exp. Ther. 262, 1044–1050, 1992. Mansour, A., Watson, S.J. and Akil, H., Opioid receptors: Past, present and future, Trends Neurosci. 18, 69–70, 1995. Yaksh, T.L. and Rudy, T.A., Studies on the direct spinal action of narcotics in the production of analgesia in the rat, J. Pharmacol. Exp. Ther. 202, 411–428, 1977. Kirby, G.C. and McQueen, D.S., Characterization of opioid receptors in the cat carotid body involved in chemosensory depression in vivo, Br. J. Pharmacol. 88, 889–898, 1986. Shook, J.E., Watkins, W.D. and Camporesi, E.M., Differential roles of opioid receptors in respiration, respiratory disease, and opiate-induced respiratory depression, Am. Rev. Respir. Dis. 142, 895–909, 1990. Tyler, D.C., Respiratory effects of pain in a child after thoracotomy, Anesthesiology 70, 873–874, 1989. Hickey, P.R., Hansen, D.D., Wessel, D.L., Lang, P., Jonas, R.A. and Elixson, E.M., Blunting of stress responses in the pulmonary circulation of infants by fentanyl, Anesth. Analg. 64, 1137–1142, 1985. Cabot, P.J., Cramond, T. and Smith, M.T., Quantitative autoradiography of peripheral opioid binding sites in rat lung, Eur. J. Pharmacol. 310, 47–53, 1996. Anjou-Lindskog, E., Broman, L., Broman, M., Holmgren, A., Settergren, G. and Ohqvist, G., Effects of intravenous anesthesia on VA/Q distribution: A study performed during ventilation with air and with 50% oxygen, supine and in the lateral position, Anesthesiology 62, 485–492, 1985. Chen, S.W., Maguire, P.A., Davies, M.F., Beatty, M.F. and Loew, G.H., Evidence for mu1-opioid receptor involvement in fentanyl-mediated respiratory depression, Eur. J. Pharmacol. 312, 241–244, 1996. Tabatabai, M., Kitahata, L.M. and Collins, J.G., Disruption of the rhythmic activity of the medullary inspiratory neurons and phrenic nerve by fentanyl and reversal with nalbuphine, Anesthesiology 70, 489–495, 1989. Weil, J.V., McCullough, R.E., Kline, J.S. and Sodal, I.E., Diminished ventilatory response to hypoxia and hypercapnia after morphine in normal man, N. Engl. J. Med. 292, 1103–1106, 1975. Bailey, P.L., Lu, J.K., Pace, N.L., Orr, J.A., White, J.L., Hamber, E.A., Slawson, M.H., Crouch, D.J. and Rollins, D.E., Effects of intrathecal morphine on the ventilatory response to hypoxia, N. Engl. J. Med. 343, 1228– 1234, 2000. Cartwright, C.R., Henson, L.C. and Ward, D.S., Effects of alfentanil on the ventilatory response to sustained hypoxia, Anesthesiology 89, 612–619, 1998. Kryger, M.H., Yacoub, O., Dosman, J., Macklem, P.T. and Anthonisen, N.R., Effect of meperidine on occlusion pressure responses to hypercapnia
Pain Management and Regional Anesthesia
104.
105. 106.
107.
108.
109. 110.
111. 112.
113.
114.
115.
116.
117.
118.
119.
497
and hypoxia with and without external inspiratory resistance, Am. Rev. Respir. Dis. 114, 333–340, 1976. Dahan, A., Sarton, E., Teppema, L. and Olievier, C., Sex-related differences in the influence of morphine on ventilatory control in humans, Anesthesiology 88, 903–913, 1998. Drummond, G.B., Comparison of decreases in ventilation caused by enflurane and fentanyl during anaesthesia, Br. J. Anaesth. 55, 825–835, 1983. Becker, L.D., Paulson, B.A., Miller, R.D., Severinghaus, J.W. and Eger, E.I., Biphasic respiratory depression after fentanyldroperidol or fentanyl alone used to supplement nitrous oxide anesthesia, Anesthesiology 44, 291–296, 1976. Bailey, P.L., Pace, N.L., Ashburn, M.A., Moll, J.W., East, K.A. and Stanley, T.H., Frequent hypoxemia and apnea after sedation with midazolam and fentanyl, Anesthesiology 73, 826–830, 1990. Looi-Lyons, L.C., Chung, F.F., Chan, V.W. and McQuestion, M., Respiratory depression: An adverse outcome during patient controlled analgesia therapy, J. Clin. Anesth. 8, 151–156, 1996. Kurth, C.D., Postoperative arterial oxygen saturation: What to expect, Anesth. Analg. 80, 1–3, 1995. Rosenberg-Adamsen, S., Kehlet, H., Dodds, C. and Rosenberg, J., Postoperative sleep disturbances: Mechanisms and clinical implications, Br. J. Anaesth. 76, 552–559, 1996. Rigg, J.R. and Rondi, P., Changes in rib cage and diaphragm contribution to ventilation after morphine, Anesthesiology 55, 507–514, 1981. Longobardo, G.S., Gothe, B., Goldman, M.D. and Cherniack, N.S., Sleep apnea considered as a control system instability, Respir. Physiol. 50, 311–333, 1982. Farney, R.J., Walker, J.M., Cloward, T.V. and Rhondeau, S., Sleepdisordered breathing associated with long-term opioid therapy, Chest 123, 632–639, 2003. Robinson, R.W., Zwillich, C.W., Bixler, E.O., Cadieux, R.J., Kales, A. and White, D.P., Effects of oral narcotics on sleep-disordered breathing in healthy adults, Chest 91, 197–203, 1987. Cronin, A.J., Keifer, J.C., Davies, M.F., King, T.S. and Bixler, E.O., Postoperative sleep disturbance: Influences of opioids and pain in humans, Sleep 24, 39–44, 2001. Berkowitz, B.A., Ngai, S.H., Yang, J.C., Hempstead, J. and Spector, S., The diposition of morphine in surgical patients, Clin. Pharmacol. Ther. 17, 629–635, 1975. Downes, J.J., Kemp, R.A. and Lambertsen, C.J., The magnitude and duration of respiratory depression due to fentanyl and meperidine in man, J. Pharmacol. Exp. Ther. 158, 416–420, 1967. Cartwright, P., Prys-Roberts, C., Gill, K., Dye, A., Stafford, M. and Gray, A., Ventilatory depression related to plasma fentanyl concentrations during and after anesthesia in humans, Anesth. Analg. 62, 966–974, 1983. Bailey, P.L., Streisand, J.B., East, K.A., East, T.D., Isern, S., Hansen, T.W., Posthuma, E.F., Rozendaal, F.W., Pace, N.L. and Stanley, T.H.,
498
120. 121.
122.
123.
124. 125. 126. 127.
128.
129.
130.
131. 132.
133. 134. 135. 136.
Bailey and Thakur Differences in magnitude and duration of opioid-induced respiratory depression and analgesia with fentanyl and sufentanil, Anesth. Analg. 70, 8–15, 1990. Peng, P.W. and Sandler, A.N., A review of the use of fentanyl analgesia in the management of acute pain in adults, Anesthesiology 90, 576–599, 1999. Bollish, S.J., Collins, C.L., Kirking, D.M. and Bartlett, R.H., Efficacy of patient-controlled versus conventional analgesia for postoperative pain, Clin. Pharm. 4, 48–52, 1985. Gourlay, G.K., Wilson, P.R. and Glynn, C.J., Pharmacodynamics and pharmacokinetics of methadone during the perioperative period, Anesthesiology 57, 458–467, 1982. Chuk, P.K., Vital signs and nurses’ choices of titrated dosages of intravenous morphine for relieving pain following cardiac surgery, J. Adv. Nurs. 30, 858–865, 1999. Rawal, N. and Wattwil, M., Respiratory depression after epidural morphine—an experimental and clinical study, Anesth. Analg. 63, 8–14, 1984. Forrest, W.H., Jr., Smethurst, P.W. and Kienitz, M.E., Self-administration of intravenous analgesics, Anesthesiology 33, 363–365, 1970. Sechzer, P.H., Studies in pain with the analgesic-demand system, Anesth. Analg. 50, 1–10, 1971. Owen, H., Szekely, S.M., Plummer, J.L., Cushnie, J.M. and Mather, L.E., Variables of patient-controlled analgesia. 2. Concurrent infusion, Anaesthesia 44, 11–13, 1989. Parker, R.K., Holtmann, B. and White, P.F., Patient-controlled analgesia. Does a concurrent opioid infusion improve pain management after surgery?, JAMA 266, 1947–1952, 1991. Fleming, B.M. and Coombs, D.W., A survey of complications documented in a quality-control analysis of patient-controlled analgesia in the postoperative patient, J. Pain Symptom Manag. 7, 463–469, 1992. McDonald, A.J. and Cooper, M.G., Patient-controlled analgesia: An appropriate method of pain control in children, Paediatr. Drugs 3, 273–284, 2001. Sandler, A., Transdermal fentanyl: Acute analgesic clinical studies, J. Pain Symptom Manag. 7, S27–S35, 1992. Bennett, R.L., Batenhorst, R.L., Bivins, B.A., Bell, R.M., Graves, D.A., Foster, T.S., Wright, B.D. and Griffen, W.O., Jr., Patient-controlled analgesia: A new concept of postoperative pain relief, Ann. Surg. 195, 700–705, 1982. McKenzie, R., Patient-controlled analgesia (PCA), Anesthesiology 69, 1027, 1988. Etches, R.C., Respiratory depression associated with patient-controlled analgesia: A review of eight cases, Can. J. Anaesth. 41, 125–132, 1994. Schug, S.A. and Torrie, J.J., Safety assessment of postoperative pain management by an acute pain service, Pain 55, 387–391, 1993. Ashburn, M.A., Love, G. and Pace, N.L., Respiratory-related critical events with intravenous patient-controlled analgesia, Clin. J. Pain 10, 52–56, 1994.
Pain Management and Regional Anesthesia
499
137. White, P.F., Mishaps with patient-controlled analgesia, Anesthesiology 66, 81– 83, 1987. 138. Baxter, A.D., Respiratory depression with patient-controlled analgesia [editorial; comment], Can. J. Anaesth. 41, 87–90, 1994. 139. Gust, R., Pecher, S., Gust, A., Hoffmann, V., Bohrer, H. and Martin, E., Effect of patient-controlled analgesia on pulmonary complications after coronary artery bypass grafting, Crit. Care Med. 27, 2218–2223, 1999. 140. Wasylak, T.J., Abbott, F.V., English, M.J. and Jeans, M.E., Reduction of postoperative morbidity following patient-controlled morphine, Can. J. Anaesth. 37, 726–731, 1990. 141. Walder, B., Schafer, M., Henzi, I. and Tramer, M.R., Efficacy and safety of patient-controlled opioid analgesia for acute postoperative pain. A quantitative systematic review, Acta Anaesthesiol. Scand. 45, 795–804, 2001. 142. Bernstein, K.Z. and Klausner, M.A., Potential dangers related to transdermal fentanyl (Duragesic) when used for postoperative pain, Dis. Colon Rectum 37, 1339–1340, 1994. 143. Nestler, E.J. and Aghajanian, G.K., Molecular and cellular basis of addiction, Science 278, 58–63, 1997. 144. Kissin, I., Brown, P.T., Robinson, C.A. and Bradley, E.L., Jr., Acute tolerance in morphine analgesia: Continuous infusion and single injection in rats, Anesthesiology 74, 166–171, 1991. 145. Teichtahl, H., Prodromidis, A., Miller, B., Cherry, G. and Kronborg, I., Sleep-disordered breathing in stable methadone programme patients: A pilot study, Addiction 96, 395–403, 2001. 146. Duttaroy, A. and Yoburn, B.C., The effect of intrinsic efficacy on opioid tolerance, Anesthesiology 82, 1226–1236, 1995. 147. Azar, I. and Turndorf, H., Severe hypertension and multiple atrial premature contractions following naloxone administration, Anesth. Analg. 58, 524–525, 1979. 148. Flacke, J.W., Flacke, W.E. and Williams, G.D., Acute pulmonary edema following naloxone reversal of high-dose morphine anesthesia, Anesthesiology 47, 376–378, 1977. 149. Kulig, K., Initial management of ingestions of toxic substances, N. Engl. J. Med. 326, 1677–1681, 1992. 150. Patschke, D., Eberlein, H.J., Hess, W., Tarnow, J. and Zimmermann, G., Antagonism of morphine with naloxone in dogs: Cardiovascular effects with special reference to the coronary circulation, Br. J. Anaesth. 49, 525–533, 1977. 151. Just, B., Delva, E., Camus, Y. and Lienhart, A., Oxygen uptake during recovery following naloxone. Relationship with intraoperative heat loss, Anesthesiology 76, 60–64, 1992. 152. Flacke, J.W., Flacke, W.E., Bloor, B.C. and Olewine, S., Effects of fentanyl, naloxone, and clonidine on hemodynamics and plasma catecholamine levels in dogs, Anesth. Analg. 62, 305–313, 1983. 153. Ngai, S.H., Berkowitz, B.A., Yang, J.C., Hempstead, J. and Spector, S., Pharmacokinetics of naloxone in rats and in man: Basis for its potency and short duration of action, Anesthesiology 44, 398–401, 1976.
500
Bailey and Thakur
154. Gan, T.J., Ginsberg, B., Glass, P.S., Fortney, J., Jhaveri, R. and Perno, R., Opioid-sparing effects of a low-dose infusion of naloxone in patientadministered morphine sulfate, Anesthesiology 87, 1075–1081, 1997. 155. Woolf, C.J., Analgesia and hyperalgesia produced in the rat by intrathecal naloxone, Brain Res. 189, 593–597, 1980. 156. Levine, J.D., Gordon, N.C. and Fields, H.L., Naloxone dose dependently produces analgesia and hyperalgesia in postoperative pain, Nature 278, 740–741, 1979. 157. Rawal, N., Schott, U., Dahlstrom, B., Inturrisi, C.E., Tandon, B., Sjostrand, U. and Wennhager, M., Influence of naloxone infusion on analgesia and respiratory depression following epidural morphine, Anesthesiology 64, 194–201, 1986. 158. Bailey, P.L., Clark, N.J., Pace, N.L., Stanley, T.H., East, K.A., van Vreeswijk, H., van de Pol, P., Clissold, M.A. and Rozendaal, W., Antagonism of postoperative opioid-induced respiratory depression: Nalbuphine versus naloxone, Anesth. Analg. 66, 1109–1114, 1987. 159. Jaffe, R.S., Moldenhauer, C.C., Hug, C.C., Jr., Finlayson, D.C., Tobia, V. and Kopel, M.E., Nalbuphine antagonism of fentanyl-induced ventilatory depression: A randomized trial, Anesthesiology 68, 254–260, 1988. 160. Latasch, L., Probst, S. and Dudziak, R., Reversal by nalbuphine of respiratory depression caused by fentanyl, Anesth. Analg. 63, 814–816, 1984. 161. Zsigmond, E.K., Durrani, Z., Barabas, E., Wang, X.Y. and Tran, L., Endocrine and hemodynamic effects of antagonism of fentanyl-induced respiratory depression by nalbuphine, Anesth. Analg. 66, 421–426, 1987. 162. Ramsay, J.G., Higgs, B.D., Wynands, J.E., Robbins, R. and Townsend, G.E., Early extubation after high-dose fentanyl anaesthesia for aortocoronary bypass surgery: Reversal of respiratory depression with low-dose nalbuphine, Can. Anaesth. Soc. J. 32, 597–606, 1985. 163. Baxter, A.D., Samson, B., Penning, J., Doran, R. and Dube, L.M., Prevention of epidural morphine-induced respiratory depression with intravenous nalbuphine infusion in post-thoracotomy patients, Can. J. Anaesth. 36, 503– 509, 1989. 164. Parker, R.K., Holtmann, B. and White, P.F., Patient-controlled epidural analgesia: Interactions between nalbuphine and hydromorphone, Anesth. Analg. 84, 757–763, 1997. 165. Joshi, G.P., Duffy, L., Chehade, J., Wesevich, J., Gajraj, N. and Johnson, E.R., Effects of prophylactic nalmefene on the incidence of morphine-related side effects in patients receiving intravenous patient-controlled analgesia, Anesthesiology 90, 1007–1011, 1999. 166. Benumof, J.L., Permanent loss of cervical spinal cord function associated with interscalene block performed under general anesthesia, Anesthesiology 93, 1541–1544, 2000. 167. Passannante, A.N., Spinal anesthesia and permanent neurologic deficit after interscalene block, Anesth. Analg. 82, 873–874, 1996. 168. Jorfeldt, L., Lofstrom, B., Pernow, B., Persson, B., Wahren, J. and Widman, B., The effect of local anaesthetics on the central circulation and respiration in man and dog, Acta Anaesthesiol. Scand. 12, 153–169, 1968.
Pain Management and Regional Anesthesia
501
169. Johnson, A. and Lofstrom, J.B., Influence of local anesthetics on ventilation, Reg. Anesth. 16, 7–12, 1991. 170. Gross, J.B., Caldwell, C.B., Shaw, L.M. and Laucks, S.O., The effect of lidocaine on the ventilatory response to carbon dioxide, Anesthesiology 59, 521–525, 1983. 171. Labaille, T., Clergue, F., Samii, K., Ecoffey, C. and Berdeaux, A., Ventilatory response to CO2 following intravenous and epidural lidocaine, Anesthesiology 63, 179–183, 1985. 172. Butterfield, N.N., Schwarz, S.K., Ries, C.R., Franciosi, L.G., Day, B. and MacLeod, B.A., Combined pre- and post-surgical bupivacaine wound infiltrations decrease opioid requirements after knee ligament reconstruction, Can. J. Anaesth. 48, 245–250, 2001. 173. McCarty, E.C., Spindler, K.P., Tingstad, E., Shyr, Y. and Higgins, M., Does intraarticular morphine improve pain control with femoral nerve block after anterior cruciate ligament reconstruction? Am. J. Sports Med. 29, 327–332, 2001. 174. Tetzlaff, J.E., Brems, J. and Dilger, J., Intraarticular morphine and bupivacaine reduces postoperative pain after rotator cuff repair, Reg. Anesth. Pain Med. 25, 611–614, 2000. 175. Vranken, J.H., Vissers, K.C., de Jongh, R. and Heylen, R., Intraarticular sufentanil administration facilitates recovery after day-case knee arthroscopy, Anesth. Analg. 92, 625–628, 2001. 176. Moiniche, S., Mikkelsen, S., Wetterslev, J. and Dahl, J.B., A qualitative systematic review of incisional local anaesthesia for postoperative pain relief after abdominal operations, Br. J. Anaesth. 81, 377–383, 1998. 177. Borgeat, A., Ekatodramis, G., Kalberer, F. and Benz, C., Acute and nonacute complications associated with interscalene block and shoulder surgery: A prospective study, Anesthesiology 95, 875–880, 2001. 178. Fackler, C.D., Perret, G.E. and Bedell, G.N., Effect of unilateral phrenic nerve section on lung function, J. Appl. Physiol. 23, 923–926, 1967. 179. Gould, L., Kaplan, S., McElhinney, A.J. and Stone, D.J., A method for the production of hemidiaphragmatic paralysis. Its application to the study of lung function in normal man, Am. Rev. Respir. Dis. 96, 812–814, 1967. 180. Urmey, W.F. and McDonald, M., Hemidiaphragmatic paresis during interscalene brachial plexus block: Effects on pulmonary function and chest wall mechanics, Anesth. Analg. 74, 352–357, 1992. 181. Sala-Blanch, X., Lazaro, J.R., Correa, J. and Gomez-Fernandez, M., Phrenic nerve block caused by interscalene brachial plexus block: Effects of digital pressure and a low volume of local anesthetic, Reg. Anesth. Pain Med. 24, 231–235, 1999. 182. Hashim, M.S. and Shevde, K., Dyspnea during interscalene block after recent coronary bypass surgery, Anesth. Analg. 89, 55–56, 1999. 183. Candido, K.D., Franco, C.D., Khan, M.A., Winnie, A.P. and Raja, D.S., Buprenorphine added to the local anesthetic for brachial plexus block to provide postoperative analgesia in outpatients, Reg. Anesth. Pain Med. 26, 352–356, 2001.
502
Bailey and Thakur
184. Casati, A., Magistris, L., Beccaria, P., Cappelleri, G., Aldegheri, G. and Fanelli, G., Improving postoperative analgesia after axillary brachial plexus anesthesia with 0.75% ropivacaine. A double-blind evaluation of adding clonidine, Minerva Anestesiol. 67, 407–412, 2001. 185. Caputo, F. and Ventura, R., Brachial plexus block. Effect of low interscalenic approach on phrenic nerve paresis, Minerva Anestesiol. 66, 195–199, 2000. 186. Mak, P.H., Irwin, M.G., Ooi, C.G. and Chow, B.F., Incidence of diaphragmatic paralysis following supraclavicular brachial plexus block and its effect on pulmonary function, Anaesthesia 56, 352–356, 2001. 187. Stadlmeyer, W., Neubauer, J., Finkl, R.O. and Groh, J., Unilateral phrenic nerve paralysis after vertical infraclavicular plexus block, Anaesthetist 49, 1030–1033, 2000. 188. Moore, D.C., Intercostal nerve block for postoperative somatic pain following surgery of thorax and upper abdomen, Br. J. Anaesth. 47 suppl, 284–286, 1975. 189. Faust, R.J. and Nauss, L.A., Post-thoracotomy intercostal block: Comparison of its effects on pulmonary function with those of intramuscular meperidine, Anesth. Analg. 55, 542–546, 1976. 190. Crawford, E.D., Skinner, D.G. and Capparell, D.B., Intercostal nerve block with thoracoabdominal incision, J. Urol. 121, 290–291, 1979. 191. Jakobson, S. and Ivarsson, I., Effects of intercostal nerve blocks (bupivacaine 0.25% and etidocaine 0.5%) on chest wall mechanics in healthy men, Acta Anaesthesiol. Scand. 21, 489–496, 1977. 192. Shanti, C.M., Carlin, A.M. and Tyburski, J.G., Incidence of pneumothorax from intercostal nerve block for analgesia in rib fractures, J. Trauma 51, 536–539, 2001. 193. Lloyd, J.W., Barnard, J.D. and Glynn, C.J., Cryoanalgesia. A new approach to pain relief, Lancet 2, 932–934, 1976. 194. Evans, P.J., Lloyd, J.W. and Green, C.J., Cryoanalgesia: The response to alterations in freeze cycle and temperature, Br. J. Anaesth. 53, 1121–1127, 1981. 195. Carter, D.C., Lee, P.W., Gill, W. and Johnston, R.J., The effect of cryosurgery on peripheral nerve function, J. R. Coll. Surg. Edinb. 17, 25–31, 1972. 196. Maiwand, M.O., Makey, A.R. and Rees, A., Cryoanalgesia after thoracotomy. Improvement of technique and review of 600 cases, J. Thorac. Cardiovasc. Surg. 92, 291–295, 1986. 197. Miguel, R. and Hubbell, D., Pain management and spirometry following thoracotomy: A prospective, randomized study of four techniques, J. Cardiothorac. Vasc. Anesth. 7, 529–534, 1993. 198. Orr, I.A., Keenan, D.J. and Dundee, J.W., Improved pain relief after thoracotomy: Use of cryoprobe and morphine infusion, Br. Med. J. (Clin. Res. Ed.) 283, 945–948, 1981. 199. Pastor, J., Morales, P., Cases, E., Cordero, P., Piqueras, A., Galan, G. and Paris, F., Evaluation of intercostal cryoanalgesia versus conventional analgesia in postthoracotomy pain, Respiration 63, 241–245, 1996.
Pain Management and Regional Anesthesia
503
200. Moorjani, N., Zhao, F., Tian, Y., Liang, C., Kaluba, J. and Maiwand, M.O., Effects of cryoanalgesia on post-thoracotomy pain and on the structure of intercostal nerves: A human prospective randomized trial and a histological study, Eur. J. Cardiothorac. Surg. 20, 502–507, 2001. 201. Kambam, J.R., Hammon, J., Parris, W.C. and Lupinetti, F.M., Intrapleural analgesia for post-thoracotomy pain and blood levels of bupivacaine following intrapleural injection, Can. J. Anaesth. 36, 106–109, 1989. 202. Kaiser, A.M., Zollinger, A., De Lorenzi, D., Largiader, F. and Weder, W., Prospective, randomized comparison of extrapleural versus epidural analgesia for postthoracotomy pain, Ann. Thorac. Surg. 66, 367–372, 1998. 203. Elman, A., Debaene, B., Magny-Metrot, C. and Murciano, G., Interpleural analgesia with bupivacaine following thoracotomy: Ineffective results of a controlled study and pharmacokinetics, J. Clin. Anesth. 5, 118–121, 1993. 204. Rosenberg, P.H., Scheinin, B.M., Lepantalo, M.J. and Lindfors, O., Continuous intrapleural infusion of bupivacaine for analgesia after thoracotomy, Anesthesiology 67, 811–813, 1987. 205. Brismar, B., Pettersson, N., Tokics, L., Strandberg, A. and Hedenstierna, G., Postoperative analgesia with intrapleural administration of bupivacaine– adrenaline, Acta Anaesthesiol. Scand. 31, 515–520, 1987. 206. Frenette, L., Boudreault, D. and Guay, J., Interpleural analgesia improves pulmonary function after cholecystectomy, Can. J. Anaesth. 38, 71–74, 1991. 207. Eng, J. and Sabanathan, S., Continuous extrapleural intercostal nerve block and post-thoracotomy pulmonary complications, Scand. J. Thorac. Cardiovasc. Surg. 26, 219–223, 1992. 208. Scheinin, B., Lindgren, L. and Rosenberg, P.H., Treatment of postthoracotomy pain with intermittent instillations of intrapleural bupivacaine, Acta Anaesthesiol. Scand. 33, 156–159, 1989. 209. Sabanathan, S., Mearns, A.J., Bickford Smith, P.J., Eng, J., Berrisford, R.G., Bibby, S.R. and Majid, M.R., Efficacy of continuous extrapleural intercostal nerve block on post-thoracotomy pain and pulmonary mechanics, Br. J. Surg. 77, 221–225, 1990. 210. Orliaguet, G. and Carli, P., Intrapleural analgesia, Ann. Fr. Anesth. Reanim. 13, 233–247, 1994. 211. Seltzer, J.L., Larijani, G.E., Goldberg, M.E. and Marr, A.T., Intrapleural bupivacaine—a kinetic and dynamic evaluation, Anesthesiology 67, 798–800, 1987. 212. Kavanagh, B.P., Katz, J. and Sandler, A.N., Pain control after thoracic surgery. A review of current techniques, Anesthesiology 81, 737–759, 1994. 213. Sabanathan, S., Smith, P.J., Pradhan, G.N., Hashimi, H., Eng, J.B. and Mearns, A.J., Continuous intercostal nerve block for pain relief after thoracotomy, Ann. Thorac. Surg. 46, 425–426, 1988. 214. Richardson, J., Sabanathan, S., Jones, J., Shah, R.D., Cheema, S. and Mearns, A.J., A prospective, randomized comparison of preoperative and continuous balanced epidural or paravertebral bupivacaine on postthoracotomy pain, pulmonary function and stress responses, Br. J. Anaesth. 83, 387–392, 1999.
504
Bailey and Thakur
215. Dhole, S., Mehta, Y., Saxena, H., Juneja, R. and Trehan, N., Comparison of continuous thoracic epidural and paravertebral blocks for postoperative analgesia after minimally invasive direct coronary artery bypass surgery, J. Cardiothorac. Vasc. Anesth. 15, 288–292, 2001. 216. Warner, D.O., Warner, M.A. and Ritman, E.L., Human chest wall function during epidural anesthesia, Anesthesiology 85, 761–773, 1996. 217. Takasaki, M. and Takahashi, T., Respiratory function during cervical and thoracic extradural analgesia in patients with normal lungs, Br. J. Anaesth. 52, 1271–1276, 1980. 218. Freund, F.G., Bonica, J.J., Ward, R.J., Akamatsu, T.J. and Kennedy, W.F. Jr., Ventilatory reserve and level of motor block during high spinal and epidural anesthesia, Anesthesiology 28, 834–837, 1967. 219. Sundberg, A., Wattwil, M. and Arvill, A., Respiratory effects of high thoracic epidural anaesthesia, Acta Anaesthesiol. Scand. 30, 215–217, 1986. 220. Stevens, R.A., Frey, K., Sheikh, T., Kao, T.C., Mikat-Stevens, M. and Morales, M., Time course of the effects of cervical epidural anesthesia on pulmonary function, Reg. Anesth. Pain Med. 23, 20–24, 1998. 221. Groeben, H., Schwalen, A., Irsfeld, S., Tarnow, J., Lipfert, P. and Hopf, H.B., High thoracic epidural anesthesia does not alter airway resistance and attenuates the response to an inhalational provocation test in patients with bronchial hyperreactivity, Anesthesiology 81, 868–874, 1994. 222. Steinbrook, R.A., Concepcion, M. and Topulos, G.P., Ventilatory responses to hypercapnia during bupivacaine spinal anesthesia, Anesth. Analg. 67, 247–252, 1988. 223. Steinbrook, R.A. and Concepcion, M., Respiratory effects of spinal anesthesia: Resting ventilation and single-breath CO2 response, Anesth. Analg. 72, 182–186, 1991. 224. Sakura, S., Saito, Y. and Kosaka, Y., Effect of extradural anaesthesia on the ventilatory response to hypoxaemia, Anaesthesia 48, 205– 209, 1993. 225. Sakura, S., Saito, Y. and Kosaka, Y., The effects of epidural anesthesia on ventilatory response to hypercapnia and hypoxia in elderly patients, Anesth. Analg. 82, 306–311, 1996. 226. Lundh, R., Hedenstierna, G. and Johansson, H., Ventilation–perfusion relationships during epidural analgesia, Acta Anaesthesiol. Scand. 27, 410–416, 1983. 227. McCarthy, G.S., The effect of thoracic extradural analgesia on pulmonary gas distribution, functional residual capacity and airway closure, Br. J. Anaesth. 48, 243–248, 1976. 228. Wahba, W.M., Craig, D.B., Don, H.F. and Becklake, M.R., The cardiorespiratory effects of thoracic epidural anaesthesia, Can. Anaesth. Soc. J. 19, 8–19, 1972. 229. Yamakage, M., Namiki, A., Tsuchida, H. and Iwasaki, H., Changes in ventilatory pattern and arterial oxygen saturation during spinal anaesthesia in man, Acta Anaesthesiol. Scand. 36, 569–571, 1992. 230. Capdevila, X., Biboulet, P., Rubenovitch, J., Serre-Cousine, O., Peray, P., Deschodt, J. and d’Athis, F., The effects of cervical epidural anesthesia with
Pain Management and Regional Anesthesia
231.
232.
233. 234.
235.
236.
237.
238.
239.
240.
241. 242.
243.
505
bupivacaine on pulmonary function in conscious patients, Anesth. Analg. 86, 1033–1038, 1998. Cuschieri, R.J., Morran, C.G., Howie, J.C. and McArdle, C.S., Postoperative pain and pulmonary complications: Comparison of three analgesic regimens, Br. J. Surg. 72, 495–498, 1985. Stenseth, R., Bjella, L., Berg, E.M., Christensen, O., Levang, O.W. and Gisvold, S.E., Effects of thoracic epidural analgesia on pulmonary function after coronary artery bypass surgery, Eur. J. Cardiothorac. Surg. 10, 859–865, 1996. Sjogren, S. and Wright, B., Respiratory changes during continuous epidural blockade, Acta Anaesthesiol. Scand. Suppl 46, 27–49, 1972. Pansard, J.L., Mankikian, B., Bertrand, M., Kieffer, E., Clergue, F. and Viars, P., Effects of thoracic extradural block on diaphragmatic electrical activity and contractility after upper abdominal surgery, Anesthesiology 78, 63–71, 1993. Conacher, I.D., Paes, M.L., Jacobson, L., Phillips, P.D. and Heaviside, D.W., Epidural analgesia following thoracic surgery. A review of two years’ experience, Anaesthesia 38, 546–551, 1983. Ross, R.A., Clarke, J.E. and Armitage, E.N., Postoperative pain prevention by continuous epidural infusion. A study of the clinical effects and the plasma concentrations obtained, Anaesthesia 35, 663–668, 1980. Callesen, T., Schouenborg, L., Nielsen, D., Guldager, H. and Kehlet, H., Combined epidural-spinal opioid-free anaesthesia and analgesia for hysterectomy, Br. J. Anaesth. 82, 881–885, 1999. Dahl, J.B., Rosenberg, J., Hansen, B.L., Hjortso, N.C. and Kehlet, H., Differential analgesic effects of low-dose epidural morphine and morphine– bupivacaine at rest and during mobilization after major abdominal surgery, Anesth. Analg. 74, 362–365, 1992. Scott, D.A., Blake, D., Buckland, M., Etches, R., Halliwell, R., Marsland, C., Merridew, G., Murphy, D., Paech, M., Schug, S.A., Turner, G., Walker, S., Huizar, K. and Gustafsson, U., A comparison of epidural ropivacaine infusion alone and in combination with 1, 2, and 4 microg/mL fentanyl for seventy-two hours of postoperative analgesia after major abdominal surgery, Anesth. Analg. 88, 857–864, 1999. Kafer, E.R., Brown, J.T., Scott, D., Findlay, J.W., Butz, R.F., Teeple, E. and Ghia, J.N., Biphasic depression of ventilatory responses to CO2 following epidural morphine, Anesthesiology 58, 418–427, 1983. Nordberg, G., Hedner, T., Mellstrand, T. and Borg, L., Pharmacokinetics of epidural morphine in man, Eur. J. Clin. Pharmacol. 26, 233–237, 1984. Camporesi, E.M., Nielsen, C.H., Bromage, P.R., Durant, P.A., Ventilatory CO2 sensitivity after intravenous and epidural morphine in volunteers, Anesth. Analg. 62, 633–640, 1983. Sandler, A.N., Stringer, D., Panos, L., Badner, N., Friedlander, M., Koren, G., Katz, J. and Klein, J., A randomized, double-blind comparison of lumbar epidural and intravenous fentanyl infusions for postthoracotomy pain relief. Analgesic, pharmacokinetic, and respiratory effects, Anesthesiology 77, 626–634, 1992.
506
Bailey and Thakur
244. Gourlay, G.K., Murphy, T.M., Plummer, J.L., Kowalski, S.R., Cherry, D.A. and Cousins, M.J., Pharmacokinetics of fentanyl in lumbar and cervical CSF following lumbar epidural and intravenous administration, Pain 38, 253–259, 1989. 245. Negre, I., Gueneron, J.P., Ecoffey, C., Penon, C., Gross, J.B., Levron, J.C. and Samii, K., Ventilatory response to carbon dioxide after intramuscular and epidural fentanyl, Anesth. Analg. 66, 707–710, 1987. 246. Renaud, B., Brichant, J.F., Clergue, F., Chauvin, M., Levron, J.C. and Viars, P., Ventilatory effects of continuous epidural infusion of fentanyl, Anesth. Analg. 67, 971–975, 1988. 247. Hansdottir, V., Bake, B. and Nordberg, G., The analgesic efficacy and adverse effects of continuous epidural sufentanil and bupivacaine infusion after thoracotomy, Anesth. Analg. 83, 394–400, 1996. 248. Rawal, N., Arner, S., Gustafsson, L.L. and Allvin, R., Present state of extradural and intrathecal opioid analgesia in Sweden. A nationwide follow-up survey, Br. J. Anaesth. 59, 791–799, 1987. 249. Stenseth, R., Sellevold, O. and Breivik, H., Epidural morphine for postoperative pain: Experience with 1085 patients, Acta Anaesthesiol. Scand. 29, 148–156, 1985. 250. Di Chiro, G., Observations on the circulation of the cerebrospinal fluid, Acta Radiol. Diagn. (Stockh.) 5, 988–1002, 1966. 251. Nordberg, G., Hedner, T., Mellstrand, T. and Borg, L., Pharmacokinetics of epidural morphine in man, Eur. J. Clin. Pharmacol. 26, 233–237, 1984. 252. Whiting, W.C., Sandler, A.N., Lau, L.C., Chovaz, P.M., Slavchenko, P., Daley, D. and Koren, G. Analgesic and respiratory effects of epidural sufentanil in patients following thoracotomy, Anesthesiology 69, 36–43, 1988. 253. Cousins, M.J. and Mather, L.E., Intrathecal and epidural administration of opioids, Anesthesiology 61, 276–310, 1984. 254. Noble, D.W., Morrison, L.M., Brockway, M.S. and McClure, J.H., Respiratory depression after extradural fentanyl, Br. J. Anaesth. 72, 251–252, 1994. 255. Etches, R.C., Sandler, A.N. and Daley, M.D., Respiratory depression and spinal opioids, Can. J. Anaesth. 36, 165–185, 1989. 256. Sakura, S., Saito, Y. and Kosaka, Y., Effect of lumbar epidural anesthesia on ventilatory response to hypercapnia in young and elderly patients, J. Clin. Anesth. 5, 109–113, 1993. 257. Modalen, A.O., Westman, L., Arlander, E., Eriksson, L.I. and Lindahl, S.G., Hypercarbic and hypoxic ventilatory responses after intrathecal administration of bupivacaine and sameridine, Anesth. Analg. 96, 570–575, 2003. 258. Bailey, P.L., Rhondeau, S., Schafer, P.G., Lu, J.K., Timmins, B.S., Foster, W., Pace, N.L. and Stanley, T.H., Dose-response pharmacology of intrathecal morphine in human volunteers (see comments), Anesthesiology 79, 49–59, 1993. 259. Lu, J.K., Manullang, T.R., Staples, M.H., Kem, S.E. and Bailey, P.L., Maternal respiratory arrests, severe hypotension, and fetal distress after administration of intrathecal, sufentanil, and bupivacaine after intravenous fentanyl, Anesthesiology 87, 170–172, 1997.
Pain Management and Regional Anesthesia
507
260. Lu, J.K., Schafer, P.G., Gardner, T.L., Pace, N.L., Zhang, J., Niu, S., Stanley, T.H. and Bailey, P.L., The dose-response pharmacology of intrathecal sufentanil in female volunteers, Anesth. Analg. 85, 372–379, 1997. 261. Hansdottir, V., Hedner, T., Woestenborghs, R. and Nordberg, G., The CSF and plasma pharmacokinetics of sufentanil after intrathecal administration, Anesthesiology 74, 264–269, 1991. 262. Lehmann, K.A., Gerhard, A., Horrichs-Haermeyer, G., Grond, S. and Zech, D., Postoperative patient-controlled analgesia with sufentanil: Analgesic efficacy and minimum effective concentrations, Acta Anaesthesiol. Scand. 35, 221–226, 1991. 263. Ummenhofer, W.C., Arends, R.H., Shen, D.D. and Bernards, C.M., Comparative spinal distribution and clearance kinetics of intrathecally administered morphine, fentanyl, alfentanil, and sufentanil, Anesthesiology 92, 739–753, 2000. 264. Shafer, S.L., Eisenach, J.C., Hood, D.D. and Tong, C., Cerebrospinal fluid pharmacokinetics and pharmacodynamics of intrathecal neostigmine methylsulfate in humans, Anesthesiology 89, 1074–1088, 1998. 265. Rawal, N., Arner, S., Gustafsson, L.L. and Allvin, R., Present state of extradural and intrathecal opioid analgesia in Sweden. A nationwide follow-up survey, Br. J. Anaesth. 59, 791–799, 1987. 266. Palmer, C.M., Emerson, S., Volgoropolous, D. and Alves, D., Doseresponse relationship of intrathecal morphine for postcesarean analgesia, Anesthesiology 90, 437–444, 1999. 267. Gwirtz, K.H., Young, J.V., Byers, R.S., Alley, C., Levin, K., Walker, S.G. and Stoelting, R.K., The safety and efficacy of intrathecal opioid analgesia for acute postoperative pain: Seven years’ experience with 5969 surgical patients at Indiana University Hospital, Anesth. Analg. 88, 599–604, 1999. 268. Boezaart, A.P., Eksteen, J.A., Spuy, G.V., Rossouw, P. and Knipe, M., Intrathecal morphine. Double-blind evaluation of optimal dosage for analgesia after major lumbar spinal surgery, Spine 24, 1131–1137, 1999. 269. Mason, N., Gondret, R., Junca, A. and Bonnet, F., Intrathecal sufentanil and morphine for post-thoracotomy pain relief, Br. J. Anaesth. 86, 236–240, 2001. 270. Kaukinen, L., Kaukinen, S., Eerola, R. and Eerola, M., The antagonistic effect of pentazocine on fentanyl induced respiratory depression compared with nalorphine and naloxone, Ann. Clin. Res. 13, 396–401, 1981. 271. Beaver, W.T., Wallenstein, S.L., Houde, R.W. and Rogers, A., A comparison of the analgesic effects of pentazocine and morphine in patients with cancer, Clin. Pharmacol. Ther. 7, 740–751, 1966. 272. Boas, R.A. and Villiger, J.W., Clinical actions of fentanyl and buprenorphine. The significance of receptor binding, Br. J. Anaesth. 57, 192–196, 1985. 273. Pedersen, J.E., Chraemmer-Jorgensen, B., Schnidt, J.F. and Risbo, A., Naloxone—a strong analgesic in combination with high-dose buprenorphine, Br. J. Anaesth. 57, 1045–1046, 1985. 274. Reynaud, M., Tracqui, A., Petit, G., Potard, D. and Courty, P., Six deaths linked to misuse of buprenorphine–benzodiazepine combinations, Am. J. Psychiatry 155, 448–449, 1998.
508
Bailey and Thakur
275. De Souza, E.B., Schmidt, W.K. and Kuhar, M.J., Nalbuphine: An autoradiographic opioid receptor binding profile in the central nervous system of an agonist/antagonist analgesic, J. Pharmacol. Exp. Ther. 244, 391–402, 1988. 276. Lee, C.R., McTavish, D. and Sorkin, E.M., Tramadol. A preliminary review of its pharmacodynamic and pharmacokinetic properties, and therapeutic potential in acute and chronic pain states, Drugs 46, 313–340, 1993. 277. Raffa, R.B., Friderichs, E., Reimann, W., Shank, R.P., Codd, E.E. and Vaught, J.L., Opioid and nonopioid components independently contribute to the mechanism of action of tramadol, an ‘atypical’ opioid analgesic, J. Pharmacol. Exp. Ther. 260, 275–285, 1992. 278. Bloch, M.B., Dyer, R.A., Heijke, S.A. and James, M.F., Tramadol infusion for postthoracotomy pain relief: A placebo-controlled comparison with epidural morphine, Anesth. Analg. 94, 523–528, 2002. 279. Hennies, H.H., Friderichs, E. and Schneider, J., Receptor binding, analgesic and antitussive potency of tramadol and other selected opioids, Arzneimittelforschung 38, 877–880, 1988. 280. Nieuwenhuijs, D., Bruce, J., Drummond, G.B., Warren, P.M. and Dahan, A., Influence of oral tramadol on the dynamic ventilatory response to carbon dioxide in healthy volunteers, Br. J. Anaesth. 87, 860–865, 2001. 281. Tarkkila, P., Tuominen, M. and Lindgren, L., Comparison of respiratory effects of tramadol and pethidine, Eur. J. Anaesthesiol. 15, 64–68, 1998. 282. Warren, P.M., Taylor, J.H., Nicholson, K.E., Wraith, P.K. and Drummond, G.B., Influence of tramadol on the ventilatory response to hypoxia in humans, Br. J. Anaesth. 85, 211–216, 2000. 283. Teppema, L.J., Nieuwenhuijs, D., Olievier, C.N. and Dahan, A., Respiratory depression by tramadol in the cat: Involvement of opioid receptors, Anesthesiology 98, 420–427, 2003. 284. Naguib, M., Seraj, M., Attia, M., Samarkandi, A.H., Seet, M. and Jaroudi, R., Perioperative antinociceptive effects of tramadol. A prospective, randomized, double-blind comparison with morphine, Can. J. Anaesth. 45, 1168–1175, 1998. 285. Gunduz, M., Ozcengiz, D., Ozbek, H. and Isik, G., A comparison of single dose caudal tramadol, tramadol plus bupivacaine and bupivacaine administration for postoperative analgesia in children, Paediatr. Anaesth. 11, 323–326, 2001. 286. Kapral, S., Gollmann, G., Waltl, B., Likar, R., Sladen, R.N., Weinstabl, C. and Lehofer, F., Tramadol added to mepivacaine prolongs the duration of an axillary brachial plexus blockade, Anesth. Analg. 88, 853– 856, 1999. 287. Murthy, B.V., Pandya, K.S., Booker, P.D., Murray, A., Lintz, W. and Terlinden, R., Pharmacokinetics of tramadol in children after i.v. or caudal epidural administration, Br. J. Anaesth. 84, 346–349, 2000. 288. Eisenach, J.C., De Kock, M. and Klimscha, W., Alpha(2)-adrenergic agonists for regional anesthesia. A clinical review of clonidine (1984–1995), Anesthesiology 85, 655–674, 1996.
Pain Management and Regional Anesthesia
509
289. Kamibayashi, T. and Maze, M., Clinical uses of alpha2-adrenergic agonists, Anesthesiology 93, 1345–1349, 2000. 290. Gordh, T., Jr., Post, C. and Olsson, Y., Evaluation of the toxicity of subarachnoid clonidine, guanfacine, and a substance P-antagonist on rat spinal cord and nerve roots: Light and electron microscopic observations after chronic intrathecal administration, Anesth. Analg. 65, 1303–1311, 1986. 291. Celly, C.S., McDonell, W.N. and Black, W.D., Cardiopulmonary effects of the alpha2-adrenoceptor agonists medetomidine and ST-91 in anesthetized sheep, J. Pharmacol. Exp. Ther. 289, 712–720, 1999. 292. Bailey, P.L., Sperry, R.J., Johnson, G.K., Eldredge, S.J., East, K.A., East, T.D., Pace, N.L. and Stanley, T.H., Respiratory effects of clonidine alone and combined with morphine, in humans, Anesthesiology 74, 43–48, 1991. 293. Jarvis, D.A., Duncan, S.R., Segal, I.S. and Maze, M., Ventilatory effects of clonidine alone and in the presence of alfentanil, in human volunteers, Anesthesiology 76, 899–905, 1992. 294. Belleville, J.P., Ward, D.S., Bloor, B.C. and Maze, M., Effects of intravenous dexmedetomidine in humans. I. Sedation, ventilation, and metabolic rate, Anesthesiology 77, 1125–1133, 1992. 295. Khan, Z.P., Ferguson, C.N. and Jones, R.M., alpha-2 and imidazoline receptor agonists. Their pharmacology and therapeutic role, Anaesthesia 54, 146–165, 1999. 296. Ooi, R., Pattison, J. and Feldman, S.A., The effects of intravenous clonidine on ventilation, Anaesthesia 46, 632–633, 1991. 297. Venn, R.M., Hell, J. and Grounds, R.M., Respiratory effects of dexmedetomidine in the surgical patient requiring intensive care, Crit. Care 4, 302–308, 2000. 298. Penon, C., Ecoffey, C. and Cohen, S.E., Ventilatory response to carbon dioxide after epidural clonidine injection, Anesth. Analg. 72, 761–764, 1991. 299. Nguyen, D., Abdul-Rasool, I., Ward, D., Hsieh, J., Kobayashi, D., Hadlock, S., Singer, F. and Bloor, B., Ventilatory effects of dexmedetomidine, atipamezole, and isoflurane in dogs, Anesthesiology 76, 573–579, 1992. 300. Narchi, P., Benhamou, D., Hamza, J. and Bouaziz, H., Ventilatory effects of epidural clonidine during the first 3 hours after caesarean section, Acta Anaesthesiol. Scand. 36, 791–795, 1992. 301. Eisenach, J., Detweiler, D. and Hood, D., Hemodynamic and analgesic actions of epidurally administered clonidine, Anesthesiology 78, 277–287, 1993. 302. Viggiano, M., Badetti, C., Roux, F., Mendizabal, H., Bernini, V. and Manelli, J.C., Controlled analgesia in a burn patient: Fentanyl sparing effect of clonidine, Ann. Fr. Anesth. Reanim. 17, 19–26, 1998. 303. Dobrydnjov, I., Axelsson, K., Thorn, S.E., Matthiesen, P., Klockhoff, H., Holmstrom, B. and Gupta, A., Clonidine combined with small-dose bupivacaine during spinal anesthesia for inguinal herniorrhaphy: A randomized double-blinded study, Anesth. Analg. 96, 1496–1503, 2003. 304. Reuben, S.S., Steinberg, R.B., Madabhushi, L. and Rosenthal, E., Intravenous regional clonidine in the management of sympathetically maintained pain, Anesthesiology 89, 527–530, 1998.
510
Bailey and Thakur
305. Gentili, M., Enel, D., Szymskiewicz, O., Mansour, F. and Bonnet, F., Postoperative analgesia by intraarticular clonidine and neostigmine in patients undergoing knee arthroscopy, Reg. Anesth. Pain Med. 26, 342–347, 2001. 306. De Negri, P., Ivani, G., Visconti, C., De Vivo, P. and Lonnqvist, P.A., The dose-response relationship for clonidine added to a postoperative continuous epidural infusion of ropivacaine in children, Anesth. Analg. 93, 71–76, 2001. 307. Gautier, P.E., De Kock, M., Fanard, L., Van Steenberge, A. and Hody, J.L., Intrathecal clonidine combined with sufentanil for labor analgesia, Anesthesiology 88, 651–656, 1998. 308. De Kock, M., Gautier, P., Fanard, L., Hody, J.L. and Lavand’homme, P., Intrathecal ropivacaine and clonidine for ambulatory knee arthroscopy: A dose-response study, Anesthesiology 94, 574–578, 2001. 309. Filos, K.S., Goudas, L.C., Patroni, O. and Polyzou, V., Hemodynamic and analgesic profile after intrathecal clonidine in humans. A dose-response study, Anesthesiology 81, 591–601, 1994. 310. Armand, S., Langlade, A., Boutros, A., Lobjoit, K., Monrigal, C., Ramboatiana, R., Rauss, A. and Bonnet, F., Meta-analysis of the efficacy of extradural clonidine to relieve postoperative pain: An impossible task, Br. J. Anaesth. 81, 126–134, 1998. 311. Woolf, C.J. and Thompson, S.W., The induction and maintenance of central sensitization is dependent on N-methyl-D-aspartic acid receptor activation; implications for the treatment of post-injury pain hypersensitivity states, Pain 44, 293–299, 1991. 312. Helmy, S.A. and Bali, A., The effect of the preemptive use of the NMDA receptor antagonist dextromethorphan on postoperative analgesic requirements, Anesth. Analg. 92, 739–744, 2001. 313. Yamamoto, T., Shimoyama, N. and Mizuguchi, T., The effects of morphine, MK-801, an NMDA antagonist, and CP-96,345, an NK1 antagonist, on the hyperesthesia evoked by carageenan injection in the rat paw, Anesthesiology 78, 124–133, 1993. 314. Finck, A.D. and Ngai, S.H., Opiate receptor mediation of ketamine analgesia, Anesthesiology 56, 291–297, 1982. 315. Collins, J.G., Effects of ketamine on low intensity tactile sensory input are not dependent upon a spinal site of action, Anesth. Analg. 65, 1123–1129, 1986. 316. Mortero, R.F., Clark, L.D., Tolan, M.M., Metz, R.J., Tsueda, K., Sheppard, R.A., The effects of small-dose ketamine on propofol sedation: Respiration, postoperative mood, perception, cognition, and pain, Anesth. Analg. 92, 1465– 1469, 2001. 317. Soliman, M.G., Brindle, G.F. and Kuster, G., Response to hypercapnia under ketamine anaesthesia, Can. Anaesth. Soc. J. 22, 486–494, 1975. 318. Stanley, V., Hunt, J., Willis, K.W. and Stephen, C.R., Cardiovascular and respiratory function with CI-581, Anesth. Analg. 47, 760–768, 1968. 319. Sarma, V.J., Use of ketamine in acute severe asthma, Acta Anaesthesiol. Scand. 36, 106–107, 1992. 320. Menigaux, C., Guignard, B., Fletcher, D., Sessler, D.I., Dupont, X. and Chauvin, M., Intraoperative small-dose ketamine enhances analgesia after outpatient knee arthroscopy, Anesth. Analg. 93, 606–612, 2001.
Pain Management and Regional Anesthesia
511
321. De Kock, M., Lavand’homme, P. and Waterloos, H., ’Balanced analgesia’ in the perioperative period: Is there a place for ketamine? Pain 92, 373–380, 2001. 322. Reeves, M., Lindholm, D.E., Myles, P.S., Fletcher, H. and Hunt, J.O., Adding ketamine to morphine for patient-controlled analgesia after major abdominal surgery: A double-blinded, randomized controlled trial, Anesth. Analg. 93, 116–120, 2001. 323. Subramaniam, K., Subramaniam, B., Pawar, D.K. and Kumar, L., Evaluation of the safety and efficacy of epidural ketamine combined with morphine for postoperative analgesia after major upper abdominal surgery, J. Clin. Anesth. 13, 339–344, 2001. 324. Kathirvel, S., Sadhasivam, S., Saxena, A., Kannan, T.R. and Ganjoo, P., Effects of intrathecal ketamine added to bupivacaine for spinal anaesthesia, Anaesthesia 55, 899–904, 2000. 325. Power, I. and Barratt, S., Analgesic agents for the postoperative period. Nonopioids, Surg. Clin. North Am. 79, 275–295, 1999. 326. Carney, D.E., Nicolette, L.A., Ratner, M.H., Minerd, A. and Baesl, T.J., Ketorolac reduces postoperative narcotic requirements, J. Pediatr. Surg. 36, 76–79, 2001. 327. Nakayama, M., Ichinose, H., Nakabayashi, K., Satoh, O., Yamamoto, S. and Namiki, A., Analgesic effect of epidural neostigmine after abdominal hysterectomy, J. Clin. Anesth. 13, 86–89, 2001. 328. Lauretti, G.R., Reis, M.P., Prado, W.A. and Klamt, J.G., Dose-response study of intrathecal morphine versus intrathecal neostigmine, their combination, or placebo for postoperative analgesia in patients undergoing anterior and posterior vaginoplasty, Anesth. Analg. 82, 1182–1187, 1996. 329. Carp, H., Jayaram, A. and Morrow, D., Intrathecal cholinergic agonists lessen bupivacaine spinal-block-induced hypotension in rats, Anesth. Analg. 79, 112– 116, 1994. 330. Kissin, I., Preemptive analgesia, Anesthesiology 93, 1138–1143, 2000. 331. Gottschalk, A., Smith, D.S., Jobes, D.R., Kennedy, S.K., Lally, S.E., Noble, V.E., Grugan, K.F., Seifert, H.A., Cheung, A., Malkowicz, S.B., Gutsche, B.B. and Wein, A.J., Preemptive epidural analgesia and recovery from radical prostatectomy: A randomized controlled trial, JAMA 279, 1076–1082, 1998. 332. Torda, T.A., Hann, P., Mills. G., De Leon, G. and Penman, D., Comparison of extradural fentanyl, bupivacaine and two fentanyl-bupivacaine mixtures for pain relief after abdominal surgery, Br. J. Anaesth. 74, 35–40, 1995. 333. Cook, T.M. and Riley, R.H., Analgesia following thoracotomy: A survey of Australian practice, Anaesth. Intensive Care 25, 520–524, 1997. 334. Park, W.Y., Thompson, J.S. and Lee, K.K., Effect of epidural anesthesia and analgesia on perioperative outcome: A randomized, controlled Veterans Affairs cooperative study, Ann. Surg. 234, 560–569, 2001. 335. Von Dossow, V., Welte, M., Zaune, U., Martin, E., Walter, M., Ruckert, J., Kox, W.J. and Spies, C.D., Thoracic epidural anesthesia combined with general anesthesia: The preferred anesthetic technique for thoracic surgery, Anesth. Analg. 92, 848–854, 2001. 336. Norris, E.J., Beattie, C., Perler, B.A., Martinez, E.A., Meinert, C.L., Anderson, G.F., Grass, J.A., Sakima, N.T., Gorman, R., Achuff, S.C.,
512
337.
338.
339.
340. 341.
342.
343. 344.
345.
346.
347.
348.
349.
Bailey and Thakur Martin, B.K., Minken, S.L., Williams, G.M. and Traystman, R.J., Doublemasked randomized trial comparing alternate combinations of intraoperative anesthesia and postoperative analgesia in abdominal aortic surgery, Anesthesiology 95, 1054–1067, 2001. Shenkman, Z., Shir, Y., Weiss, Y.G., Bleiberg, B. and Gross, D., The effects of cardiac surgery on early and late pulmonary functions, Acta Anaesthesiol. Scand. 41, 1193–1199, 1997. van Belle, A.F., Wesseling, G.J., Penn, O.C. and Wouters, E.F., Postoperative pulmonary function abnormalities after coronary artery bypass surgery, Respir. Med. 86, 195–199, 1992. Chaney, M.A., Smith, K.R., Barclay, J.C. and Slogoff, S., Large-dose intrathecal morphine for coronary artery bypass grafting, Anesth. Analg. 83, 215–222, 1996. Mathews, E.T. and Abrams, L.D., Intrathecal morphine in open heart surgery, Lancet 2, 543, 1980. Taylor, A., Healy, M., McCarroll, M. and Moriarty, D.C., Intrathecal morphine: One year’s experience in cardiac surgical patients, J. Cardiothorac. Vasc. Anesth. 10, 225–228, 1996. Loick, H.M., Schmidt, C., Van Aken, H., Junker, R., Erren, M., Berendes, E., Rolf, N., Meissner, A., Schmid, C., Scheld, H.H. and Mollhoff, T., High thoracic epidural anesthesia, but not clonidine, attenuates the perioperative stress response via sympatholysis and reduces the release of troponin T in patients undergoing coronary artery bypass grafting, Anesth. Analg. 88, 701–709, 1999. Gravlee, G.P., Epidural analgesia and coronary artery bypass grafting: The controversy continues, J. Cardiothorac. Vasc. Anesth. 17, 151–153, 2003. Scott, N.B., Turfrey, D.J., Ray, D.A., Nzewi, O., Sutcliffe, N.P., Lal, A.B., Norrie, J., Nagels, W.J. and Ramayya, G.P., A prospective randomized study of the potential benefits of thoracic epidural anesthesia and analgesia in patients undergoing coronary artery bypass grafting, Anesth. Analg. 93, 528–535, 2001. Turfrey, D.J., Ray, D.A., Sutcliffe, N.P., Ramayya, P., Kenny, G.N. and Scott, N.B., Thoracic epidural anaesthesia for coronary artery bypass graft surgery. Effects on postoperative complications, Anaesthesia 52, 1090–1095, 1997. Pastor, M.C., Sanchez, M.J., Casas, M.A., Mateu, J. and Bataller, M.L., Thoracic epidural analgesia in coronary artery bypass graft surgery: Seven years’ experience, J. Cardiothorac. Vasc. Anesth. 17, 154–159, 2003. Liem, T.H., Hasenbos, M.A., Booij, L.H. and Gielen, M.J., Coronary artery bypass grafting using two different anesthetic techniques: Part 2: Postoperative outcome, J. Cardiothorac. Vasc. Anesth. 6, 156–161, 1992. Priestley, M.C., Cope, L., Halliwell, R., Gibson, P., Chard, R.B., Skinner, M. and Klineberg, P.L., Thoracic epidural anesthesia for cardiac surgery: The effects on tracheal intubation time and length of hospital stay, Anesth. Analg. 94, 275–282, 2002. Pflug, A.E. and Bonica, J.J., Physiopathology and control of postoperative pain, Arch. Surg. 112, 773–781, 1977.
14 Ventilatory Effects of Medications Used for Moderate and Deep Sedation
JEFFREY B. GROSS
DANTE A. CERZA
University of Connecticut School of Medicine Farmington, Connecticut
University of Pennsylvania School of Medicine Philadelphia, Pennsylvania
I.
Introduction
In 1971, Monheim coined the term sedalgesia to describe a drug-induced alteration of consciousness which would lessen the impact of an unpleasant experience upon a conscious patient during uncomfortable dental procedures [1]. The following year, Lee used the term conscious sedation to describe a state of modified consciousness which allowed patients to be comfortable while maintaining their own cardiorespiratory function [2]. For over a century, inhalation agents such as nitrous oxide had been used for this purpose. By the mid-1970s, intravenous conscious sedation with benzodiazepines, barbiturates, and major tranquilizers increased in popularity because it did not require the use of specialized gas delivery systems [3]. In addition, the use of conscious sedation spread from the dental office to many medical venues where patients were subjected to uncomfortable procedures such as upper and lower GI endoscopy, bronchoscopy, and cystoscopy. Pediatric procedures, especially those that are not particularly uncomfortable such as radiologic studies, also proved suitable to conscious sedation. 513
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In 1986, midazolam was introduced and quickly supplanted diazepam for most procedural sedation. Advantages of midazolam included the fact that injection did not cause local pain or venous irritation, and that it seemed to provide better amnesia than diazepam. Within two years, however, it became apparent that conscious sedation with midazolam entailed significant risks. In fact, the United States Food and Drug Administration considered limiting the use of midazolam after its use for conscious sedation resulted in at least 40 reported deaths [4]. Many of these were related to respiratory depression, resulting either from midazolam alone or from the combination of midazolam and an opioid. As a result of these events, the manufacturer changed the labeling to reduce the recommended dose and emphasize the need for careful patient monitoring during conscious sedation. In 1994, the American Society of Anesthesiologists established a task force to develop guidelines for conscious sedation by nonanesthesiologists. This task force introduced the term sedation and analgesia (reminiscent of Monheim’s sedalgesia [1]) to describe the state wherein patients were adequately sedated to undergo uncomfortable procedures but still able to maintain their own vital functions. The resulting guidelines provide recommendations regarding drug administration and patient monitoring. Since most unfavorable outcomes from sedation and analgesia result from respiratory compromise, most of the recommendations are designed to: (a) reduce the risk of respiratory depression; (b) increase the likelihood that it will be detected in a timely fashion (by monitoring pulmonary ventilation and arterial oxygenation), and (c) maximize the likelihood that patients who inadvertently become overly sedated will be successfully rescued [5]. In 1999, in recognition of the fact that sedation and analgesia represents a continuum, the American Society of Anesthesiologists adopted its Continuum of Depth of Sedation (Table 14.1) [6]. This document includes functional definitions of minimal sedation (anxiolysis), moderate sedation (conscious sedation), deep sedation, and general anesthesia. Ideally, during conscious sedation no interventions are required to maintain a patent airway, and spontaneous ventilation is adequate. However, as discussed below, many of the medications administered for conscious sedation have the potential to decrease airway patency and/or depress spontaneous ventilatory drive. As a result, practitioners administering these drugs for conscious sedation must be able to recognize these consequences and manage them appropriately. While anesthesiologists routinely care for patients with obstructed airways and depressed ventilation, caring for these patients may present particular challenges for non-anesthesiologists who administer conscious sedation. For this reason, it is particularly important that such individuals understand the potential
Responsiveness Airway Spontaneous Ventilation Cardiovascular Function
Minimal Sedation (Anxiolysis)
Moderate Sedation/Analgesia (Conscious Sedation)
Deep Sedation/Analgesia
General Anesthesia
Normal response to verbal stimulation Unaffected Unaffected
Purposefula response to verbal or tactile stimulation No intervention required Adequate
Purposefula response following repeated or painful stimulation Intervention may be required May be inadequate
Unarousable, even with painful stimulus Intervention often required Frequently inadequate
Unaffected
Usually maintained
Usually maintained
May be impaired
Minimal Sedation is a drug-induced state during which patients respond normally to verbal commands. Although cognitive function and coordination may be impaired, ventilatory and cardiovascular functions are unaffected. Moderate Sedation/Analgesia (Conscious Sedation) is a drug-induced depression of consciousness during which patients respond purposefullya to verbal commands, either alone or accompanied by light tactile stimulation. No interventions are required to maintain a patent airway, and spontaneous ventilation is adequate. Cardiovascular function is usually maintained. Deep Sedation/Analgesia is a drug-induced depression of consciousness during which patients cannot be easily aroused, but respond purposefullya following repeated or painful stimulation. The ability to independently maintain ventilatory function may be impaired. Patients may require assistance in maintaining a patent airway, and spontaneous ventilation may be inadequate. Cardiovascular function is usually maintained. General Anesthesia is a drug-induced loss of consciousness during which patients are not arousable, even by painful stimulation. The ability to independently maintain ventilatory function is often impaired. Patients often require assistance in maintaining a patent airway, and positive pressure ventilation may be required because of depressed spontaneous ventilation or drug-induced depression of neuromuscular function. Cardiovascular function may be impaired. Source: Data from Ref. 6. a Reflex withdrawal from a painful stimulus is NOT considered a purposeful response.
Ventilatory Effects of Medications Used for Sedation
Table 14.1 Definitions of Levels of Sedation and Analgesia
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ventilatory depressant effects of medications commonly administered during moderate (conscious) sedation. In 2001, the American Society of Anesthesiologists updated its Guidelines for Sedation and Analgesia by Non-Anesthesiologists [6]. The revision specifically addressed issues related to both moderate and deep sedation as described in the 1999 definitions. Additionally, because of their increasing popularity among non-anesthesiologists, the updated guidelines provide recommendations for the safe use of rapidly acting sedative/hypnotic agents such as propofol and methohexital to provide moderate or deep sedation. Most patients undergoing moderate or deep sedation for medical procedures require both a sedative and an opioid analgesic. The sedative medication reduces the patient’s anxiety and awareness of the procedure, and often produces antegrade amnesia. In addition to potentiating the relaxation provided by sedative drugs, opioids reduce the patient’s response to painful interventions. If opioids are not used during painful interventions, a much deeper level of sedation may be required to keep the patient comfortable and cooperative. In the discussion that follows, we will address, in turn, the ventilatory effects of the sedatives, opioids, and adjuvant drugs, and their combinations, which are commonly used during moderate or deep sedation. II.
Sedatives
A. Benzodiazepines Diazepam
Diazepam is a hydrophobic benzodiazepine that can be administered orally, intramuscularly, or intravenously to provide sedation and amnesia. Diazepam is frequently administered orally, in a dose of 0.1–0.2 mg kg1, for the management of insomnia, anxiety, and muscle spasms, as well as for minimal sedation prior to medical or dental procedures. These patients almost never develop clinically significant respiratory depression unless other sedative medications or alcohol are taken concomitantly. In fact, the package insert for ValiumÕ tablets does not even mention respiratory depression as a possible adverse reaction at the recommended dose [7]. This is consistent with the observations of Delpierre et al., who found that in adult patients oral diazepam, 20 mg per day, had no effect on the slope of the CO2 ventilatory response curve. They did, however observe a modest decrease in the unstimulated P0.1 (airway pressure 100 ms after the start of an occluded breath) following diazepam; this was attributed to the skeletal muscle relaxant effects of diazepam rather than to a depressant effect on ventilatory drive [8]. Similarly, Mak et al. observed that oral diazepam 0.08 mg kg1 did not affect the ventilatory response to CO2 as measured
Ventilatory Effects of Medications Used for Sedation
517
by the Read rebreathing method, despite the observation that their subjects were clinically sedated [9]. Using rebreathing techniques, other investigators were able to demonstrate effects of oral diazepam on ventilatory control. Gilmartin et al. found that the slope of the ventilatory response to CO2 decreased by 40% (P 5 0.01), and that the slope of the P0.1 response to CO2 decreased by 61% (P 5 0.01) in adult volunteers given 10 mg of oral diazepam [10]. Utting and Pleuvry demonstrated that oral diazepam 5 mg had no effect on the slope of the CO2 response but caused a statistically significant 3 mm Hg rightward displacement of the curve [11]. Rapoport et al. found that 10 mg of diazepam administered orally caused V_ E at PETCO2 ¼ 55 mm Hg to decrease by 28% (P 5 0.01); changes in the slopes of the V_ E or P0.1 vs. PCO2 curves were not significant [12]. Beaupre et al. determined that 10 mg of oral diazepam had no effect on the ventilatory response to CO2 and a 28% decrease in the P0.1 response to CO2 in patients with chronic obstructive pulmonary disease [13]. The discrepancies among these results may be related to the details of the study protocols. The volunteers in the Gilmartin [10] and Rapoport [12] studies received 10 mg of diazepam, and post-diazepam measurements were made within an hour of diazepam administration. In contrast, Mak et al. only administered 5 mg of diazepam, which may be an insufficient dose to cause significant respiratory depression [9]. The patients in Delpierre’s study received diazepam over a course of two days before their postdiazepam ventilatory drive determinations, which were conducted several hours after the last dose of diazepam. Thus, they may have developed tolerance to the ventilatory effect of the sedative [8]. Two additional studies evaluated the effect of oral diazepam on the upper airway musculature. Leiter et al. [14] demonstrated that 10 mg of oral diazepam not only depressed the ventilatory response to CO2 but also decreased genioglossal muscle EMG activity. This effect was most pronounced in subjects over 30 years of age, and was greater than the simultaneously measured depression in tidal volume, suggesting that airway obstruction is likely to make a greater contribution to hypoventilation in these individuals. In contrast, Philip-Joe¨t et al. [15] were unable to demonstrate a consistent effect of oral diazepam 0.15 mg kg1 on upper airway collapsibility, as measured by the critical airway opening pressure. There are at least 15 published investigations regarding the effect of parenteral diazepam on ventilatory control (Table 14.2). When administered intramuscularly, diazepam appears to modestly depress ventilatory drive. With very low doses (0.02–0.06 mg kg1), Steen et al. [16] found no significant effect on ventilatory drive; with higher doses, Gasser et al. [21] found a modest shift in the CO2 response and Cegla [20] observed an inconsistent decrease in its slope. Lakshminarayan et al. [22] found no
518
Table 14.2
Studies Evaluating the Effects of Diazepam on Ventilatory Drive
Author
N
Dose/Route
6
Design
15 13
Catchlove (1971) [19]
10
0.11 mg kg1 i.v.
Cegla (1973) [20]
10
10–20 mg i.m.
CO2 rebreathe
Gasser (1976) [21]
5
7.5–15 mg i.m.
CO2 rebreathe
Lakshminarayan (1976) [22] Jordan (1980) [23]
8
10 mg i.m.
6
15 mg i.v.
Forster (1980) [24]
8
0.3 mg kg1 i.v.
CO2 rebreathe; Hypoxic rebreathe Resting values; CO2 rebreathe CO2 rebreathe
Clergue (1981) [25]
10
0.14 mg kg1 i.v.
CO2 rebreathe
!VE vs. CO2 shift
Resting values Resting values; CO2 rebreathe Resting values; CO2 rebreathe
"PCO2; #V_ E ; #TV "PCO2; #TV !V_ E vs. CO2 slope "PCO2; #V_ E ; #TV; !V_ E vs. CO2 slope #V_ E vs. CO2 slope @ 10 mg; !V_ E vs. CO2 slope @ 20 mg !V_ E vs. CO2 slope (2–3 mm Hg right shift of V_ E vs. CO2) !V_ E vs. CO2 response slope; #V_ E response to hypoxia "PCO2; #V_ E ; #V_ E vs. CO2 slope #V_ E vs. CO2 slope; #P0.1 vs. CO2 slope #V_ E ; #V_ E vs. CO2 slope; #P0.1 vs. CO2 slope
Resting values; CO2 rebreathe
Comment – – Variable effects on CO2 response Patients with COPD; Variable effects on CO2 response Higher baseline slope in 10 mg group Peak effect at 1–2 h
Peak effect at 30 min Partial reversal with naloxone 15 mg – –
Gross and Cerza
Dalen (1969) [17] Catchlove (1971) [18]
0.02–0.06 mg kg1 i.m. 5–10 mg i.v. 0.14 mg kg1 i.v.
Steen (1966) [16]
Outcome
8
0.4 mg kg1 i.v.
Power (1983) [27] Bourke (1984) [28]
7 6
Spaulding (1984) [29] Bailey (1986) [30]
0.15 mg kg1 i.v. 0.29 mg kg1 i.v.
!V_ E vs. CO2 slope !Hypercapnic V_ E
10
0.4 mg kg1 i.v.
CO2 rebreathe
#V_ E vs. CO2 slope
24
0.1 mg kg1 i.v.
CO2 rebreathe
!V_ E vs. CO2 slope; #P0.1 vs. CO2 slope
8
Resting V_ E ; Resting PCO2 Resting PCO2 Hypoxic rebreathe
!V_ E ; "PCO2
Resting PCO2; Resting SpO2
!PCO2; #SpO2 from 1–80 min
Sunzel (1988) [32] Mora (1989) [33]
8 10
0.15 mg kg1 i.v. 3 0.45 mg kg1 i.v. 1.6 mg kg1 i.v.
Zakko (1999) [34]
49
0.11 mg kg1 i.v.
Berggren (1987) [31]
#V_ E vs. CO2 slope
Dual-isohypercapnic CO2 response CO2 rebreathe Isohypercapnia (PCO2 ¼ 46 mm Hg)
Year of publication appears in parentheses. " Indicates an increase in the indicated variable. # Indicates a decrease in the indicated variable. ! Indicates no change in the indicated variable.
"PCO2 #V_ E response to hypoxia
Significant depression 1–25 min after injection Statistical significance 49% decrease in V_ E did not reach statistical significance Further decrease in slope with physostigmine Significant between-subject variability; P50.051 for V_ E vs. CO2 slope #Tidal volume; "Resting rate EC50 ¼ 423 ng ml1 Diazepam infusion during clinical procedures Patients also received meperidine 0.58 mg kg1
Ventilatory Effects of Medications Used for Sedation
Gross (1982) [26]
519
520
Gross and Cerza
change in CO2 response but a significant decrease in the ‘shape factor’ of the hypoxic response. While most studies of intravenous diazepam reveal a significant decrease in one or more measures of ventilatory control, there are some interesting exceptions. Although Power et al. demonstrated a 25% decrease in the slope of the ventilatory response to CO2 following diazepam 0.15 mg kg1 i.v., they could not demonstrate statistical significance, most likely because their six-subject study was underpowered [27]. Similarly, Bourke et al. found a 49% decrease in V_ E at PETCO2 ¼ 46 mm Hg following diazepam 0.29 mg kg1 i.v.; again, because they studied only six subjects they were unable to demonstrate statistical significance [28]. Bailey et al. found a significant decrease in the slope of the P0.1 ventilatory response to CO2 following diazepam 0.1 mg kg1 i.v. [30]. However, because of significant intersubject variability, they were unable to demonstrate a significant depression of the ventilatory response; the fact that P 5 0.051 suggests the possibility of a type II error. Interestingly, the hypoxic ventilatory drive in Mora’s [33] patients decreased by only 22% despite the fact that they received a huge dose (1.6 mg kg1) of diazepam. Most likely, this was related to the fact that diazepam was administered in 2.5–5.0 mg increments over the course of a clinical procedure which typically lasted 1–2 h. This suggests that respiratory depression following incrementally administered diazepam may reach a plateau. In fact, this effect was observed by Sunzel et al. [32], who estimated that the EC50 for diazepam-induced ventilatory depression was 423 ng ml1. In conclusion, it appears that the effect of diazepam on ventilatory drive depends to some extent upon the route of administration. Oral diazepam in doses typically administered for anxiolysis may modestly depress the ventilatory response to CO2. The clinical observation that patients seldom, if ever develop respiratory compromise as a result, may be related to the relatively gradual onset of ventilatory depression following orally administered diazepam. Under these circumstances, the gradually developing decrease in V_ E causes a gradual increase in PaCO2, which mitigates the effect of ventilatory depression on resting ventilation [35,36]. Similarly, intramuscular diazepam takes effect gradually, with the peak ventilatory effect occurring between 30 and 120 min after injection [21,22]. While this may reduce the likelihood of untoward clinical consequences, resting values of PCO2 and V_ E may be affected [17]. When administered intravenously, diazepam is more likely to produce significant ventilatory depression. This is likely to be related to the more rapid increase in the concentration of diazepam in the ventilatory control centers: the peak ventilatory effect of i.v. diazepam occurs approximately 3 min after injection, with gradual recovery beginning about 20 min later [26]. Thus, in the clinical situation, the ventilatory depression caused by i.v. diazepam is less likely to be offset by a simultaneous rise in PaCO2.
Ventilatory Effects of Medications Used for Sedation
521
Midazolam
The effect of midazolam on ventilatory control has been the subject of more than a dozen published investigations (Table 14.3). An oral midazolam preparation intended for pediatric sedation was introduced in 1998. Due to extensive first-pass metabolism, the bioavailability is less than 50%, unless a patient is concomitantly taking medications that inhibit the 3A4 isoenzyme of cytochrome P450 (e.g., erythromycin, ketoconazole). Neither Power et al. [39] nor Mak et al. [9] were able to demonstrate any effect of oral midazolam on the ventilatory response to carbon dioxide. This may be related to the relatively low doses of midazolam they studied (15 and 7.5 mg, respectively), as well as to the extensive first-pass metabolism of the drug. Clinically, Litman et al. [49] found that oral midazolam 0.7 mg kg1 minimally affected spontaneous ventilation, and did not cause upper airway obstruction. Mean PETCO2 was 42.7 mm Hg after the drug was administered, the maximum observed PETCO2 was 51 mm Hg, and mean SpO2 exceeded 99% (FiO2 5 0.3). On the basis of these data, it appears that in recommended doses, oral midazolam is not associated with a significant risk of ventilatory depression or airway obstruction. In contrast, intravenous midazolam does pose a significant risk of respiratory depression and airway obstruction. In the first published study of midazolam-induced ventilatory depression, Forster et al. [24] demonstrated a 33% decrease in the ventilatory response to CO2 following midazolam 0.15 mg kg1 i.v.; that this was a central effect was demonstrated by a 44% decrease in the P0.1 response. The effect was similar to that observed after diazepam 0.3 mg kg1, suggesting that midazolam was twice as potent as diazepam. Subsequently, Gross et al. [37] demonstrated a 34% decrease in the slope of the CO2 ventilatory response following administration of midazolam 0.2 mg kg1 to healthy volunteers; the peak depression occurred 2 min after midazolam injection, with minimal recovery during the 15 min study period. In patients with pre-existing obstructive pulmonary disease, the CO2 ventilatory response decreased by 79%, suggesting that pre-existing lung disease increases the susceptibility to midazolam-induced ventilatory depression. Once again, peak ventilatory depression occurred 2 min after midazolam injection, with minimal recovery during the 15 min observation period. Sunzel et al. [32] determined the effect of three 0.05 mg kg1 incremental doses of midazolam on PaCO2. They found that the increase in PaCO2 reached a plateau of 6 mm Hg when plasma midazolam concentrations exceed 200 ng ml1; the EC50 for midazolam-induced increase in PaCO2 was 53 ng ml1. Using an isocapnic rebreathing technique, Alexander et al. [40] demonstrated that midazolam 0.1 mg kg1 caused a 53% decrease in the slope of the hypoxic ventilatory response; physostigmine 2 mg modestly increased subjects’ level of consciousness but did not affect their hypoxic
522
Table 14.3 Studies Evaluating the Effects of Midazolam on Ventilatory Drive Author Forster (1980) [24] Gross (1983) [37]
N
Dose/Route 1
8 0.15 mg kg
i.v.
14 0.2 mg kg1 i.v.
Design
#V_ E vs. PCO2 slope; #P0.1 vs. CO2 slope Dual-isohypercapnic #V_ E vs. CO2 slope CO2 response
CO2 rebreathe
8 0.05–0.2 mg kg1 i.v.
Resting values
Power (1983) [27]
7 0.075 mg kg1 i.v.
CO2 rebreathe
Power (1984) [39] Berggren (1987) [31] Alexander (1988) [40] Sunzel (1988) [32] Bailey (1990) [41]
9 15 mg p.o. 8 0.05 mg kg1 i.v. 3
CO2 rebreathe Resting PCO2; Resting V_ E Hypoxic rebreathe
8 0.15 mg kg1 i.v. 12 0.05 mg kg1 i.v.
Dahan (1991) [42]
5
0.025 mg kg1 i.v. þ0.014 mg kg1 h1
Resting PaCO2 CO2 rebreathe; Resting SpO2 Sustained isocapnic i.v. hypoxia
#TV; "Respir. rate; "PETCO2 ; #SaO2 !V_ E vs. CO2 slope
!V_ E vs. CO2 slope "PCO2; ! VE #V_ E response to hypoxia "PaCO2 !V_ E vs. CO2 slope; No hypoxia !AHR "HVD
Comment Effect same as with diazepam 0.3 mg kg1 Effect more pronounced in patients with COPD than controls. Peak at 2 min Peak respir. depression at 2 min; Peak CO2 at 6 min 38% decrease in slope at 3 min after injection did not reach statistical significance – No difference from diazepam Physostigmine no different than placebo EC50 ¼ 53 ng ml1 Significant change when fentanyl added #Tidal volume during midazolam infusion
Gross and Cerza
Forster (1983) [38]
8 0.1 mg kg1 i.v.
Outcome
15 0.13 mg kg1 i.v.
Alexander (1992) [44] 10 0.1 mg kg1 i.v.
CO2 rebreathe; CO2 steady-state
#V_ E vs. CO2 slope; #V_ E @PCO ¼ 46 mm Hg 2
Dual-isohypercapnic #V_ E vs. CO2 slope CO2 response
Blouin (1993) [45]
12 0.12 mg kg1 i.v.
Hypoxic rebreathe
#V_ E response to hypoxia
Mak (1993) [9]
6
CO2 rebreathe; Hypoxic rebreathe
Flo¨gel (1993) [46]
32 33 mg i.v. infusion
Nagyova (1993) [47]
6
!V_ E vs. CO2 slope; !V_ E response to hypoxia #V_ E vs. CO2 slope; "V_ E @PCO2 ¼ 46 mm Hg !AHR
7.5 mg p.o.
CO2 pseudorebreathing Sustained isocapnic hypoxia CO2 rebreathe
Mora (1995) [48]
Target-controlled i.v. infusion (40 ng ml1) 16 28 mg i.v. infusion
Litman (1997) [49]
34 0.7 mg kg1 p.o.
Resting PCO2; SpO2
Gross (1996) [50]
12 Target-controlled i.v. infusion (79 ng ml1) 49 0.03 mg kg1 i.v.
Steady-state CO2; Hypoxic rebreathe Resting PCO2; Resting SpO2
Zakko (1999) [34]
!V_ E vs. CO2 slope PCO2 445 mm Hg in 34% of patients (max 51 mm Hg); !SpO2 #V_ E vs. CO2 slope; #V_ E response to hypoxia "PCO2 from 5 to 30 min #SpO2 from 1 to 80 min
"V_ E vs. CO2 slope; "V_ E at PCO2 ¼ 46 with flumazenil Maximum effect independent of injection rate Complete reversal with flumazenil No effect from flumazenil – No effect from flumazenil Mean plasma concentration 364 ng ml1 No significant change in PCO2 with 40% N2O Background alfentanil infusion 9 ng ml1 –
Ventilatory Effects of Medications Used for Sedation
Gross (1991) [43]
523
524
Gross and Cerza
ventilatory drive. Dahan et al. [42] used a dynamic end-tidal forcing technique to measure both acute hypoxic response (AHR) and hypoxic ventilatory decline (HVD). They found that a midazolam infusion (0.025 mg kg1 over 10 min followed by 0.014 mg kg1 h1) only reduced the AHR by 27% (P ¼ NS). However, during midazolam infusion, HVD was more pronounced, so the V_ E following 15 min of hypoxia was 54% lower during midazolam infusion when compared with control conditions. One possible reason for the rash of adverse outcomes shortly after the introduction of midazolam is that the dosing was inappropriate. Based on animal data, midazolam was felt to be twice as potent as diazepam. The data of Forster et al. [24] seemed to confirm this view, since respiratory depression following midazolam 0.15 mg kg1 was the same as that following diazepam 0.3 mg kg1. However, the doses of midazolam and diazepam used in the Forster study exceeded those subsequently shown to cause maximal depression of ventilatory drive by Sunzel et al. [32]. Thus, the observation that these doses caused equivalent respiratory depressant effects did not prove that midazolam is twice as potent as diazepam. The study of Zakko et al. [34] addressed this issue; using a double-blind technique, these investigators demonstrated that midazolam was 3.4 times more potent than diazepam in decreasing level of consciousness. However, at this dose ratio, midazolam was associated with more respiratory depression than diazepam (Figure 14.1); this suggests that midazolam is
42 Midazolam
CO2 Tension (mm Hg)
41
Diazepam
40 39 38 37 36 35 34 −10
0
10
20
30
40
50
60
70
80
90
Time after initial benzodiazepine (min)
Figure 14.1 End-tidal carbon dioxide tension during sedation with meperidine and either midazolam or diazepam. Initial dose of benzodiazepine and opioid was given at time ¼ 0. Midazolam values were significantly higher than baseline from 5 to 30 min after injection, whereas diazepam values were significantly higher than baseline from 65 to 70 min after injection (P 5 0.05) (Data from Ref. 34).
Ventilatory Effects of Medications Used for Sedation
525
probably more than 3.4 times as potent as diazepam with regard to respiratory depression. Can midazolam-induced ventilatory depression be mitigated by injecting the drug more slowly? Mora et al. [48] administered midazolam 0.3–0.5 mg min1 to adult patients undergoing endoscopic procedures. Despite large total midazolam doses of 0.4 mg kg1, they were unable to demonstrate significant depression of the ventilatory response to CO2 as determined by a rebreathing technique; this suggests that slow administration can, in fact, reduce the risk of ventilatory depression. In contrast, Alexander et al. [44] found that while the onset of respiratory depression was delayed, the ultimate degree of depression of the CO2 ventilatory response was independent of whether midazolam 0.1 mg kg1 was injected quickly (over 15 sec) or slowly (over 5 min) (Figure 14.2). This is consistent with the findings of Morrow et al., who observed that hypoxia (SpO2 5 90% breathing room air) was less likely when midazolam and meperidine were administered as a single bolus dose rather than by dose titration during colonoscopy [51]. The ASA Guidelines for Sedation and Analgesia by Non-Anesthesiologists recommend administering incremental
2.25 2
Slope (l·min−1·mm Hg−1)
1.75 *
1.5
*
1.25 * 1 0.75
Fast injection
0.5
Slow injection 0.25 0 0
2
4
6
8
10
12
14
16
18
20
Time (min)
Figure 14.2 Comparative slopes of the response to CO2 (l min1 mm Hg1) when midazolam 0.1 mg kg1 was injected fast (over 15 sec as indicated by the arrow) and slowly (over 5 min, as indicated by the hatched line). Values are means SE. *P 5 0.05 between injection rates at the indicated times (Data from Ref. 44).
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Gross and Cerza
doses of sedative medications; if sufficient time is allowed for each dose to reach its peak respiratory and sedative effect before the next dose is given, it may be possible to reduce the risk of excessive respiratory depression [5]. Midazolam may also affect upper airway patency. Montravers et al. [52] demonstrated a six-fold increase in supraglottic airway resistance 5 min after midazolam 0.1 mg kg1 i.v. In a subsequent study, these investigators found that midazolam 0.1 mg kg1 increased work of breathing from 3 to 6.3 J min1; this was primarily related to increased upper airway resistance during inspiration [53]. Midazolam-induced airway obstruction can be relieved by advancement of the mandible (jaw thrust) [54], nasal CPAP [55], or flumazenil [56]. Agents Used to Reverse Benzodiazepine-Induced Ventilatory Depression Flumazenil
Initial investigations into the effect of flumazenil on benzodiazepine-induced ventilatory depression were inconclusive. Mora et al. [33] found that flumazenil 1 mg did not increase hypoxic ventilatory response in patients who had received a mean diazepam dose of 0.97 mg kg1. Despite the large dose of diazepam, hypoxic drive was only decreased by about 22%, perhaps explaining the absence of a statistically significant increase following flumazenil. Carter et al. [57] found that in patients who received a mean dose of 6 mg of midazolam, oxygen saturations decreased from 94.7% to 92.7%. Although oxygen saturation increased to 94.3% following flumazenil, the increase was not statistically significant. Subsequent investigations yielded more encouraging results. Gross et al. [43] found that flumazenil 1 mg had little effect on the slope of the ventilatory response to CO2 following midazolam 0.13 mg kg1. However, the displacement of the CO2 response curve (V_ E at PETCO2 ¼ 46 mm Hg) returned to baseline within 3 min of flumazenil administration. These investigators also examined the effect of flumazenil on hypoxic ventilatory drive [45]. Midazolam decreased the slope of the hypoxic ventilatory response by 42%; following flumazenil, this variable returned to its baseline value, while it was unaffected by a placebo. Forster et al. [58] demonstrated that flumazenil 0.1 mg kg1, alone, has no ventilatory effects; in addition, this dose of flumazenil completely blocked the ventilatory depressant effect of subsequently administered midazolam 0.1 mg kg1. Flo¨gel et al. studied the effect of flumazenil (1, 3, and 10 mg) on V_ E at a fixed PETCO2 ¼ 46 mm Hg during a midazolam infusion [46], finding that all three doses completely reversed midazolam-induced ventilatory depression. Since opioids are commonly coadministered with benzodiazepines during moderate sedation, it is clinically important to know whether flumazenil can effectively reverse benzodiazepine-induced ventilatory
Ventilatory Effects of Medications Used for Sedation
527
depression in the presence of an opioid. Gross et al. [50] found that an alfentanil infusion (plasma concentration ¼ 9 ng ml1) decreased the V_ E response to CO2 from 2.14 to 1.43 l min1 mm Hg1. Midazolam infusion (plasma concentration ¼ 79 ng ml1) further reduced this response to 0.87 l min1 mm Hg1. Following flumazenil, the V_ E response increased to 1.47 l min1 mm Hg1, indicating the antagonist completely reversed that component of ventilatory depression attributable to the benzodiazepine, despite continued opioid-induced ventilatory depression. Physostigmine
Prior to the introduction of flumazenil, the centrally acting anticholinesterase drug physostigmine was sometimes administered to reverse benzodiazepineinduced sedation [59,60]. Spaulding et al. [29] found that although physostigmine increased subjects’ level of consciousness following sedation with diazepam, the ventilatory response to CO2 actually decreased when compared with a placebo; this may be related to a direct ventilatory depressant effect of acetylcholine within the CNS, as demonstrated by Gesell et al. [61]. In contrast, Bourke et al. found that in the subset of their subjects whose ventilatory drive was depressed by diazepam, physostigmine tended to increase ventilation during isohypercarbia [28]. B. Barbiturates Pentobarbital
Pentobarbital is commonly used to provide moderate sedation for patients undergoing non-invasive diagnostic procedures [62]; when it is used for painful procedures, patients may become disinhibited and uncooperative, so other regimens are preferred. When administered orally, clinically effective doses of pentobarbital (1–3 mg kg1) minimally affect ventilatory drive. Finn et al. [63] found no significant effect on the position of the CO2 response curve following 200 mg of oral pentobarbital. Similarly, Murray et al. [64] found that neither 50 nor 150 mg of pentobarbital p.o. had any effect on the position or slope of the ventilatory response to CO2. Power et al. [39] found a 25% decrease in the slope of the ventilatory response to CO2 following pentobarbital 100 mg p.o.; their results did not reach statistical significance, perhaps because with only nine subjects, their study was underpowered. Intramuscular pentobarbital seems to affect ventilatory drive. For example, Brown et al. [65] found a slight but statistically significant right shift of ventilatory response to CO2 following pentobarbital 60 and 180 mg i.m; the shifts were small (averaging less than 1 mm Hg), and the effect on the slope of the ventilatory response was not reported. Similarly, Gasser et al. [66] demonstrated a small (1–2 mm Hg) but significant rightward shift of the CO2 response curve following pentobarbital 50 and 150 mg i.m; the
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slope of the CO2 response was not significantly affected. Finally, Hirshman et al. [67] studied the effect of pentobarbital 2 mg kg1 i.m. on the ventilatory response to hypoxia, finding that neither resting V_ E nor PETCO2 was affected by the drug. However, the shape factor A of the hyperbola relating V_ E to PETO2 decreased by 35% following pentobarbital. The maximum depression was observed 1 h after the drug was administered; after another hour, hypoxic ventilatory drive had returned to baseline. Interestingly, they noted that their patients could be subdivided into two groups: in the five volunteers with the lowest oxygen uptake, the shape factor ‘A’ decreased by 53% (P 5 0.05); in the other five subjects, ‘A’ decreased by only 8% (P ¼ NS). It appears that when administered alone in clinically appropriate doses, pentobarbital minimally affects ventilatory control. However, if the dose is excessive [68], or if opioids are administered concomitantly [69], significant respiratory depression may occur. Methohexital
Methohexital is an ultrashort-acting oxybarbiturate that may be administered either rectally or by intravenous infusion to produce moderate or deep sedation. Rectal methohexital is primarily used in pediatric patients, either to provide sedation for procedures or as a premedication before the induction of general anesthesia. Audenaert et al. [69] found no change in the respiratory rate of children who received methohexital 30 mg kg1 p.r. However, Larsson et al. [70] found that rectal methohexital 20 and 30 mg kg1 increased resting PCO2 in arterialized capillary blood by 4 and 5 mm Hg, respectively; in each group, there were some patients whose PCO2 increased to more than 60 mm Hg during methohexital sedation. Daniels et al. [71] studied the effect of rectal methohexital 25–30 mg kg1 on oxygen saturation in infants and children. They observed that only 2 of 31 patients had any episodes of hypoxemia (SpO2 90%); both of these episodes appear to have been related to airway obstruction, as they occurred when patients had their heads flexed over a parent’s shoulder while awaiting their procedure. In a study of 648 patients who received rectal methohexital 30 mg kg1, Audenaert et al. [72] found that oxygen desaturation (SpO2 93%) occurred in 4%. Most of these were successfully treated with blow-by oxygen or a jaw-thrust maneuver; however two patients developed airway obstruction or laryngospasm, and required positive pressure ventilation. Intravenous methohexital significantly depresses ventilatory drive. Kay [73] studied the ventilatory depressant effect of methohexital in intubated, spontaneously breathing patients lightly anesthetized with N2O and enflurane. He demonstrated that methohexital 0.5 mg kg1 caused a 57% depression of the unstimulated P0.1 with peak respiratory depression occurring one minute after the drug was injected. Choi et al. [74] used the
Ventilatory Effects of Medications Used for Sedation
529
dual-isohypercapnic technique to study the effect of i.v. methohexital on ventilatory drive. They observed that the slope of the ventilatory response to CO2 decreased by 94% (P 5 0.05) following methohexital 1.5 mg kg1 i.v.; the V_ E response to CO2 remained significantly depressed for 8 min after injection, even though the mean time to recovery of consciousness was only 4 min. Bickler et al. [75] examined the effect of methohexital infusion on respiratory drive and ventilatory mechanics; although their patients were breathing spontaneously, they were deeply sedated, with methohexital concentrations of 6 mg ml1. They observed a 44% decrease in the slope of the V_ E response to CO2; in those individuals who breathed through a facemask, there was no change in functional residual capacity associated with methohexital infusion. Approximately 40% of the tidal volume was contributed by the rib cage; this fraction did not change during methohexital infusion. There have been several clinical investigations of the effect of methohexital on spontaneous ventilation. Wise et al. [76] administered deep sedation with intermittent doses of methohexital (mean total dose 5.7 mg min1) to patients undergoing dental restoration. Two-thirds of their 30 patients developed airway obstruction despite continuous chin elevation. Five patients developed hypoxemia (PaO2 5 70 mm Hg); three of these were visibly cyanotic and received supplemental oxygen. Mackenzie et al. [77] found that methohexital infusion at a rate of 0.2 mg kg1 min1 was associated with a 5 breath min1 increase in respiratory rate and a 3 mm Hg increase in PETCO2 . In most of the other clinical studies of methohexitalinduced sedation, patients received additional sedative or analgesic agents. For example, Allen et al. [78] sedated patients with a combination of methohexital (1 mg kg1 initial dose followed by 20–30 mg increments) and N2O (70% in O2). They observed a significant, progressive increase in PaCO2 during deep sedation, which resolved after discontinuation of the methohexital and N2O. Thiopental
Thiopental is most commonly used to provide deep sedation or for induction of general anesthesia. Therefore, most studies of the ventilatory effects of thiopental have been performed under these circumstances (Table 14.4). Rigg et al. [80] found that during sedation with a thiopental infusion (maximum blood concentration 6.2 mg ml1), the ventilatory response to CO2 by the rebreathing technique decreased by more than 50%; interestingly, resting V_ E and PETCO2 , measured immediately before the rebreathing tests, remained within 10% of their control values. Knill et al. [81] studied five subjects during thiopental infusion titrated to produce sedation rather than anesthesia. They found there was no change in resting
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Table 14.4
Studies Evaluating the Effects of Intravenous Thiopental on Ventilatory Drive
Author
N
Duffin (1976) [79]
14
Dose 0.2 mg kg1 min1
Design Change in V_ E with 2 breaths of O2 CO2 rebreathe
#V_ E (10%)
CO2 rebreathe; Hypoxic rebreathe
!V_ E vs. CO2 slope !V_ E response to hypoxia #V_ E vs. CO2 slope #V_ E response to hypoxia #V_ E ; !P0.1 #V_ E vs. CO2 slope
8
3–6 mg kg1 infusion
Knill (1978) [81]
5
Knill (1978) [81]
10
Germain (1982) [82] Gross (1983) [37]
28 14
3 mg kg1 plus 0.2 mg kg1 min1 infusion 4–6 mg kg1 plus 0.3 mg kg1 min1 infusion 7.4 mg kg1 3.5 mg kg1
Taylor (1986) [83]
36
4 mg kg1
Resting V_ E ; Resting P0.1 Dual-isohypercapnic CO2 response Resting V_ E ; Resting rate
Grounds (1987) [84]
6
4 mg kg1
Resting V_ E ; Resting rate
Valtonen (1989) [85]
15
3–4 mg kg1
Resting PETCO2 Dual-isohypercapnic CO2 response Resting V_ E ; Resting rate
8
4 mg kg1
Spens (1996) [87]
23
4 mg kg1
CO2 rebreathe; Hypoxic rebreathe
#V_ E vs. CO2 slope
#Resting V_ E ; !Resting rate #Resting V_ E ; !Resting rate !Resting PETCO2 #V_ E vs. CO2 slope #Resting V_ E ; #Resting rate
Comment No change in V_ E during halothane #Tidal volume; "Respiratory rate Subjects studied in a state of moderate sedation Subjects studied during general anesthesia (intubated) !TI/TTOT Normal and COPD patients Some patients received papaveretum and/or scopolamine All patients received papaveretum and scopolamine Patients premedicated with diazepam 0.2 mg kg1 i.v. Slope returned to baseline within 5 min Peak effect at 1 min (mean duration of apnea 22 sec)
Gross and Cerza
Rigg (1976) [80]
Blouin (1991) [86]
Outcome
Ventilatory Effects of Medications Used for Sedation
531
ventilation; the slope of the ventilatory response to CO2 decreased by 24%, but the change did not reach statistical significance (perhaps because of the small number of subjects). Knill et al. [81] also studied 10 patients who received a thiopental infusion that produced a state of general anesthesia. In these patients, the slope of the ventilatory response to CO2 was reduced by 65% (P 5 0.05). Blouin et al. [86] followed the time course of depression of the ventilatory response to CO2 in eight healthy volunteers. They found that the slope of the CO2 response decreased by 56% after thiopental 4 mg kg1. Peak ventilatory depression occurred 30 sec after thiopental injection; within 5 min after injection ventilatory response returned to baseline. Studies by Germain et al. [82], Taylor et al. [83], Grounds et al. [84], and Spens et al. [87] evaluated the effect of thiopental on resting ventilatory variables, without CO2 stimulation. They all found a significant decrease in resting V_ E , primarily because of a decrease in tidal volume rather than respiratory rate. Knill et al. [81] performed the only direct evaluation of the effect of thiopental on hypoxic ventilatory response. During thiopental sedation, they found only a 9% decrease in the shape factor A of the hypoxic ventilatory response (P ¼ NS). However, with an infusion tailored to produce light anesthesia, ‘A’ decreased by 55% (P 5 0.05). Duffin et al. [79] studied the effect of a thiopental infusion on peripheral chemoreceptor response by administering two breaths of 100% O2 to their subjects, who had been breathing room air. They found that under these circumstances, oxygen decreased resting minute ventilation by approximately 10%, indicating that at least some degree of hypoxic response was retained; this response was significantly greater than that observed during halothane anesthesia. Drummond [88] evaluated the effect of thiopental on the airway musculature. With increasing levels of sedation, there was a tendency toward decreased activity of the genioglossus muscle. In addition, activity of the scalene and strap muscles changed from tonic to phasic. If the airway became obstructed, the phasic activity increased in amplitude until the obstruction was relieved, at which time the level of phasic activity also decreased. The author concluded that a change in the pattern of muscle activation is more important than the decrease in its amplitude in causing airway obstruction during thiopental sedation. It appears that both thiopental and methohexital cause depression of the ventilatory responses to CO2 and hypoxia in proportion to the depth of sedation or anesthesia. In addition, airway obstruction becomes more likely as sedation deepens. Since patients receiving moderate sedation with barbiturates tend to become disinhibited, thiopental and methohexital are most often used to provide deep sedation or general anesthesia. Under these circumstances, significant respiratory depression and airway obstruction are likely to occur. For this reason, the 2001 update of the American
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Gross and Cerza
Society of Anesthesiologists Guidelines for Sedation and Analgesia by Non-Anesthesiologists recommends that patients receiving these medications always receive care appropriate for ‘deep sedation’ even if ‘moderate sedation’ is intended [6]. C. Propofol
When administered in doses providing deep sedation or general anesthesia, propofol significantly decreases resting ventilation, increases resting PETCO2 and decreases the ventilatory response to CO2 (Table 14.5). Mackenzie et al. [77] observed that propofol (200–300 mg kg1 min1) increased resting PETCO2 by 8 mm Hg; respiratory rate was also increased, indicating that propofol’s primary effect on ventilation is to decrease the tidal volume. The increase in PETCO2 was significantly greater than that observed during an equivalent infusion of methohexital. In lightly anesthetized patients receiving propofol 100 mg kg1 min1, Goodman et al. [89] found that the ventilatory response to CO2, as measured by the rebreathing method, decreased to 58% of the control value. Doubling the infusion rate did not further depress ventilatory drive. Allsop et al. [90] were unable to demonstrate an effect of 100 mg kg1 min1 of propofol on the slope of the CO2 response; Goodman suggested that this may have been related to type II error associated with the wide variability in Allsop’s observations [96]. Blouin et al. [86] found that the slope of the ventilatory response to CO2 decreased by 56% following a single 2.5 mg kg1 dose of propofol. Peak depression of the CO2 response curve occurred 90 sec after propofol administration, and the CO2 response remained depressed for 20 min; the ventilatory depression lasted appreciably longer than clinically assessed sedation, suggesting that propofol-induced ventilatory depression may persist despite clinical recovery (Figure 14.3). Also, ventilatory depression lasted appreciably longer than that observed following an equivalent dose (4 mg kg1) of thiopental. Even relatively low doses of propofol may significantly affect the ventilatory response to CO2. For example Vangerven et al. [91] found that when infused at 100 mg kg1 min1, propofol significantly increased resting PETCO2 . Nieuwenhuijs et al. [95] used dynamic end-tidal forcing of CO2 tension to determine whether propofol-induced ventilatory depression was primarily central or peripheral in origin. Using a targetcontrolled infusion, they found that plasma propofol concentrations of 0.5 mg ml1 (mean BIS ¼ 84; Bispectral index, Aspect Medical Systems, Newton, MA) decreased the slope of the CO2 response by 20%. This change was exclusively related to the more slowly responding central ventilatory control system. The rapidly responding peripheral chemoreceptor system was unaffected. At plasma propofol concentrations of 1.3 mg ml1
Dose
Author
N
Mackenzie (1985) [77]
40
200–300 mg kg1 min1
Resting rate; Resting PETCO2
"Resting rate; "PETCO2
Taylor (1986) [83]
38
2.5 mg kg1
Resting rate; Resting V_ E
#Resting rate; #V_ E
Grounds (1987) [84]
6
2.5 mg kg1
#V_ E ; #Inspiratory flow; !Resting rate
Goodman (1987) [89]
7
2.5 mg kg1 bolus; 100–200 mg kg1 min1 infusion 100 mg kg1 min1 infusion
Resting V_ E ; Inspiratory flow; Resting rate Resting V_ E ; Resting PCO2; CO2 rebreathe Resting V_ E ; Resting PCO2; CO2 rebreathe
1.5–2.0 mg kg1
Resting PETCO2
!PETCO2
2.5 mg kg1
Dual-isohypercapnic CO2 response
#V_ E vs. CO2 slope
Allsop (1988) [90]
19
Valtonen (1989) [85]
15
Blouin (1991) [86]
8
Design
Outcome
Comment Greater increase in PETCO2 than with equivalent dose of methohexital Some patients received papaveretum and/or scopolamine All patients received papaveretum and scopolamine
!V_ E ; #PETCO2 ; #V_ E vs. CO2 slope #V_ E ; "PETCO2 ; !V_ E vs. CO2 slope
Subjects who received papaveretum had right shift of CO2 response curve during propofol Subjects premedicated with diazepam 0.2 mg kg1 i.v. Slope remained depressed for 20 min (significantly longer than with thiopental)
533
(Continued)
Ventilatory Effects of Medications Used for Sedation
Table 14.5 Studies Evaluating the Effects of Intravenous Propofol on Ventilatory Drive
534
Table 14.5 Continued Author
N
Dose
Design
Resting PETCO2 20 1 mg kg1 bolus; 100 mg kg1 min1 infusion Hypoxic rebreathe 8 1 mg kg1bolus; 85 mg kg1 min1 infusion Nagyova (1995) [93] 12 Target controlled infusion Dynamic end-tidal (plasma concentrations 0.1, forcing of PETO2 0.5, 1 and 2 mg kg1 min1)
Vangerven (1992) [91] Blouin (1993) [92]
Spens (1996) [87]
23 2.5 mg kg1 bolus
"Resting PETCO2 #V_ E response to hypoxia # Acute V_ E response to hypoxia (dose dependent) #Resting V_ E ; #Resting rate
Comment – PETCO2 clamped to 46–48 mm Hg Significantly less depression than equipotent doses of halothane Peak depression of V_ E at 90 sec (all subjects apneic); mean duration of apnea 88 sec
Dynamic end-tidal forcing #AHR; "HVD/AHR of PETO2 ; AHR; HVD Dynamic end-tidal forcing of PETCO2
#Central sensitivity to CO2; !Peripheral sensitivity to CO2
Gross and Cerza
Nieuwenhuijs (2000) 10 Target controlled infusion [94] (plasma concentrations 0.6 mg ml1) Nieuwenhuijs (2001) 10 Target controlled [95] infusion (plasma concentrations 0.5, 1.3 mg ml1)
Resting V_ E ; Resting rate
Outcome
Ventilatory Effects of Medications Used for Sedation
535
2.5 Slope (l·min−1·mm Hg−1)
Thiopental
Propofol
2 1.5 1 0.5 *#
*+ 0 −2
0
2
4
6
8
*#
*# 10
12
14
16
18
20
Time (min)
Figure 14.3 Slope of ventilatory response to carbon dioxide (l min1 mm Hg1) following propofol 2.5 mg kg1 and thiopental 4 mg kg1 injected at time ¼ 0. *P 5 0.05 versus preinjection value for propofol. þ P 5 0.05 versus preinjection value for thiopental. #P 5 0.05 for propofol versus thiopental during the same time period (Data from Ref. 86).
(mean BIS ¼ 67), the slope of the CO2 response was 31% lower than baseline. Again, this change was entirely related to a decrease in the centrally mediated response; peripheral chemoreceptor responses were unaffected. Moderate sedation with propofol also depresses hypoxic ventilatory drive. Nagyova et al. [93] found a dose-dependent decrease in the AHR during target-controlled infusions of propofol. At plasma concentrations of 0.52 mg ml1, the AHR decreased by 22%; at plasma concentrations of 2.1 mg ml1, the depression of AHR was 61%. Interestingly, the authors found that propofol-induced depression of the AHR was significantly less than that associated with equipotent, subanesthetic doses of halothane. Blouin et al. [92] studied volunteers who were sedated with a propofol infusion to a level where their eyes closed spontaneously but they responded to loud verbal commands (plasma concentration 2.2 mg ml1); hypoxic ventilatory response, measured at PETCO2 ¼ 46–48 mm Hg, decreased by 80% under these conditions. Within 30 min of discontinuing the infusion, ventilatory response returned to baseline. Nieuwenhuijs et al. [94] determined the effect of low concentrations of propofol (0.6 mg ml1) on the AHR and on HVD. They found that compared with pre-propofol values, there was a 37% decrease in AHR during propofol infusion (mean BIS ¼ 76). HVD, expressed as a fraction of the AHR (HVD/AHR),
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Gross and Cerza
was 0.39 in control studies and 0.55 during propofol, indicating that propofol affects central as well as peripheral ventilatory control mechanisms. Like other medications used for deep sedation and general anesthesia, propofol may cause significant airway obstruction. Litman et al. [97] found that during propofol sedation in children, the narrowest part of the airway was at the level of the soft palate, rather than at the base of the tongue. During sedation the anteroposterior diameter exceeded the transverse diameter; this relationship was reversed as the patients awakened. Hammer et al. [98] found that a jaw-thrust maneuver was effective in relieving propofol-induced airway obstruction in infants; this intervention increased minute ventilation by 61% during sedation with intermittent propofol 1–2 mg kg1. Fiberoptic examination revealed that the jaw thrust caused an anterior displacement of the epiglottis, enlarging the laryngeal inlet. Also, patients sedated with propofol are less likely to develop laryngospasm than those receiving thiopental. McKeating et al. [99] found that following propofol 2.5 mg kg1, 82% of patients had immobile vocal cords; the comparable figure following thiopental 4 mg kg1 was only 22%. Due to its propensity to cause significant depression of the ventilatory responses to both CO2 and hypoxia, pulmonary ventilation of patients receiving propofol sedation should be continually monitored [6]. For example, when Vargo et al. [100] sedated patients undergoing endoscopic procedures with a propofol infusion, they found that 6 of 10 patients had one or more periods of clinically significant apnea. Of note is the fact that apnea always preceded hypoxemia, as detected by pulse oximetry. The authors emphasize that a second clinician, trained in monitoring and life support techniques, must continuously monitor the patient, and intervene appropriately if ventilation, oxygenation, or cardiovascular stability are compromised. D. Ketamine
The ventilatory effects of ketamine are critically dependent upon the dose and rate of administration of the drug. Low doses of ketamine may cause activation of the sympathetic nervous system and concomitant stimulation of ventilation. Morel et al. [101] found that ketamine 1 mg kg1 administered over 5 min caused a 66% increase in V_ E , which was evident from 5 to 20 min after the start of drug infusion; both respiratory rate and tidal volume were increased. Bourke et al. [102] found that cumulative doses of ketamine of less than 0.7 mg kg1 minimally affected V_ E during sustained hypercapnia (PETCO2 ¼ 50 mm Hg). Higher doses of ketamine (up to 3 mg kg1) caused a significant decrease in the CO2-stimulated V_ E , indicating a right-shift of the CO2 response curve; however, the slope of the ventilatory response CO2 was unaffected.
Ventilatory Effects of Medications Used for Sedation
537
The relationship between ketamine dose and its ventilatory effects was clearly demonstrated by Hama et al. [103]. They found that a 2 mg kg1 i.v. dose of ketamine decreased the slope of the V_ E response to CO2 by 39%. The decrease in ventilation was accompanied by a decrease in respiratory rate rather than tidal volume. In contrast, during a subsequent ketamine infusion (40 mg kg1 min1), the slope of the V_ E response to CO2 returned to its baseline value. However, minute ventilation at PETCO2 ¼ 60 mm Hg was depressed by 22% (P 5 0.05). This suggests that high concentrations of ketamine depress CO2 responsiveness, while lower doses merely shift the curve to the right. Since the drug is absorbed more slowly, intramuscular administration of ketamine might be expected to present a decreased risk of respiratory depression. However, Mitchell et al. [104] and Smith et al. [105] each report a case where respiratory arrest occurred following intramuscular injection of ketamine 4 mg kg1. E.
Droperidol
Droperidol is a butyrophenone neuroleptic agent, which is typically administered in combination with an opioid to provide moderate or deep sedation. In this section, we will discuss the ventilatory effects of droperidol alone; the effects of droperidol–opioid combinations will be addressed in the section dedicated to drug combinations (see below). Droperidol minimally affects resting ventilation; Soroker et al. [106] found that droperidol 5 mg i.m. caused a slight (8%) but statistically significant (P 5 0.05) decrease in resting minute ventilation with no change in resting PaO2 or PaCO2. Lehmann et al. [107] found that droperidol 5 mg i.v. caused a 28% increase in the slope of the V_ E response to CO2 rebreathing, but the change did not achieve statistical significance because of variability between subjects. Prokocimer et al. [108] studied the effects of a larger dose of droperidol, 0.3 mg kg1 i.v. Overall, they found no significant change in the slope of the V_ E or P0.1 responses to a rebreathing CO2 challenge. However, one of their subjects demonstrated a three-fold increase in the V_ E response, while another exhibited a 62% decrease in this response; the authors suggest that the wide variability in responses may have resulted from side effects such as anxiety or dyskinesia which occurred during their study. In contrast, droperidol appears to consistently increase hypoxic ventilatory response. Ward [109] found that 2.5 mg of droperidol i.v. caused a 90% increase in the slope of the hypoxic ventilatory response as assessed by dynamic end-tidal forcing. Furthermore, this study demonstrated that droperidol effectively blocks dopamine-induced depression of the hypoxic ventilatory response, suggesting that blockade of endogenous dopamine may partially account for the droperidol-induced stimulation of hypoxic ventilatory response.
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Gross and Cerza
By itself, droperidol minimally affects the ventilatory response to CO2 and augments the ventilatory response to hypoxemia. This suggests that use of this drug for moderate or deep sedation does not increase the risk of respiratory depression. However, the associated dysphoria, long duration of action, and concerns about electrocardiographic QT interval prolongation have markedly reduced the popularity of droperidol–opioid combinations for most procedures. F.
Diphenhydramine
Diphenhydramine is an H1-receptor antihistamine widely used in overthe-counter anti-anxiety and sleep medications. Clinically, it is also used to provide anxiolysis and moderate sedation as well as to treat opioid-related side effects such as nausea and pruritus. Alexander et al. [110] found that diphenhydramine 0.7 mg kg1 did not affect the ventilatory response to CO2 during hyperoxia (i.e., with hypoxic drive completely suppressed). Similarly, there was no effect on the hypoxic ventilatory response as measured under normocarbic conditions (PETCO2 ¼ 46 mm Hg). However, when the hypoxic ventilatory response was measured under hypercarbic conditions (PETCO2 ¼ 54 mm Hg), there was a significant increase following administration of diphenhydramine. This interaction can be illustrated by the effect of diphenhydramine on the Oxford Fans, which show the CO2 ventilatory response at different arterial oxygen saturations (Figure 14.4). Since diphenhydramine is frequently administered concomitantly with an opioid, Babenco et al. [111] administered a low-dose alfentanil infusion, which decreased the slope of the CO2 response curve by 27% (P 5 0.05). Following diphenhydramine 0.7 mg kg1, the slope increased significantly (P 5 0.05) to slightly above its baseline value, although the curve did remain slightly shifted to the right. This suggests that the ventilatory stimulation of diphenhydramine partially counteracts the ventilatory depression associated with m-opioid agonists such as alfentanil, and that patients receiving diphenhydramine with an opioid are at no greater risk for respiratory depression than from the opioid alone (see below). G. Chloral Hydrate
Chloral hydrate is commonly used to provide sedation for infants and children undergoing diagnostic and dental restorative procedures. Lees et al. [112] studied 13 infants (mean gestational age 41 weeks) first during natural sleep, and again following chloral hydrate 50 mg kg1 p.o. They found that the V_ E response to CO2, as determined by the steady-state method, was 38% higher during chloral hydrate sedation than during natural sleep (P ¼ NS).
Ventilatory Effects of Medications Used for Sedation
539
100 Pre-diphenhydramine
Minute ventilation (l/min)
90
SpO
80
2
70
70%
60
80%
50 90%
40
100%
30 20 10 0 30
35
40
45
50
55
60
PCO (mm Hg) 2
100 70%
Minute ventilation (l/min)
90 80
Post-diphenhydramine 80%
70 60
90%
50 40
100%
30 20 10 0 30
35
40
45
50
55
60
PCO (mm Hg) 2
Figure 14.4 Oxford fans showing ventilatory response to CO2 at different oxygen saturations before and after diphenhydramine 0.7 mg kg1 (Adapted from Ref. 110).
Resting ventilation was 20% higher following chloral hydrate (P 5 0.05). There was an associated 28% increase in O2 consumption compared with natural sleep (P 5 0.01). In contrast, Litman et al. [113] found evidence of respiratory depression in children (1–9 years old) receiving chloral hydrate 70 mg kg1 alone or in combination with N2O. While breathing room air after taking chloral hydrate, 77% of the subjects hypoventilated
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(PETCO2 4 45 mm Hg). With the addition of 30% N2O, the fraction of children hypoventilating increased to 94%. Hypoxia (SpO2 5 92%) did not occur, but it should be noted that during N2O administration, FiO2 was 0.3. The tendency of hypoventilation to worsen when N2O is administered to patients sedated with chloral hydrate is emphasized by Cote´’s review of adverse events during pediatric sedation [114]. These investigators found that addition of N2O to other sedating medications such as chloral hydrate was frequently associated with adverse outcomes. Chloral hydrate also may predispose patients to upper airway obstruction. In animal studies, Hershenson et al. [115] found that peak genioglossus EMG activity decreased by 63% (P 5 0.001) after nasogastric administration of chloral hydrate. Conversely, diaphragmatic EMG activity was not significantly depressed following chloral hydrate. These data, taken together with those presented above, suggest the airway obstruction may pose a greater risk than ventilatory depression per se in infants and children given chloral hydrate alone. III.
Opioids
The ventilatory depressant and central analgesic effects of the opioids depend primarily upon their activity at m opioid receptors. Classically, the receptors responsible for analgesia were thought to be indistinguishable from those responsible for ventilatory depression. More recent investigation suggests that there are two subtypes: m1 producing analgesia and m2 producing respiratory depression [130]. Unfortunately, at the present time there are no clinically available opioid agonists that are specific for the m1 receptor. Hence, increasing levels of opioid-induced analgesia are invariably linked to increasing degrees of ventilatory depression. A. Morphine
As the prototypical m-opioid receptor agonist, the respiratory effects of morphine have been the most thoroughly investigated. Since the pharmacodynamic effects of the other m-agonists are very similar, morphine provides a good standard of comparison and will be considered first. As shown in Table 14.6, morphine consistently reduces resting ventilation and increases resting PCO2. It also decreases the ventilatory response to carbon dioxide. However, the nature of this effect appears to depend upon both the dose of morphine and the measurement technique. When the CO2 response is determined by a steady-state technique, low doses of morphine (0.05–0.30 mg kg1) appear to cause a right-shift of the CO2 response without a significant change in its slope [120,124,136]. This is similar to the effect of natural sleep, which produces a modest right-shift of the CO2 response without change in slope [137]. However, if the dose is
Studies Evaluating the Effects of Morphine on Ventilatory Drive N
Dose/Route
Design
Outcome
Comment
Campbell (1964) [116] Weil (1975) [117]
6 6
0.14–0.28 mgkg1 i.v. 7.5 mg s.c.
Single CO2 challenge CO2 rebreathe; Hypoxic rebreathe
– –
Knill (1976) [118]
5
0.1 mg kg1 i.m.
CO2 rebreathe
Rigg (1978) [119] Gal (1982) [120]
17 6
0.15 mg kg1 i.m. 0.15–0.60 mg kg1 i.v.
CO2 rebreathe CO2 rebreathe
Camporesi (1983) [121] Arunasalam (1983) [122]
10 26
10 mg i.v. 0.14 mg kg1 i.v.
CO2 rebreathe Resting V_ E , Resting PCO2, Resting rate
Daykin (1986) [123]
24
0.14 mg kg1 i.v.
CO2 rebreathe; Resting PCO2
"PaCO2 # V_ E vs. CO2 slope; # V_ E response to hypoxia # V_ E vs. CO2 slope; Right shift P0.1 vs. CO2 # V_ E vs. CO2 slope Right shift of V_ E vs. CO2 at 0.15 mg kg1; # V_ E vs. CO2 slope at 0.6 mg kg1 # V_ E vs. CO2 slope # Resting V_ E ; " Resting PCO2; #Resting rate # V_ E vs. CO2 slope; "Resting PCO2
Bourke (1989) [124]
4
0.07–0.21 mg kg1 i.v.
CO2 steady-state; CO2 rebreathe
Author
Right-shift of steadystate V_ E vs. CO2; # V_ E vs. CO2 slope during rebreathe
– – –
Resolved after 3 h No difference between young (28–37 yr) and old (65–82 yrs) patients No difference between young (18–29 yr) and old (66–85 yr) patients Discrepancy related to increased resting PCO2 after morphine & decreased initial CO2 step during rebreathing measurements
541
(Continued)
Ventilatory Effects of Medications Used for Sedation
Table 14.6
542
Table 14.6
Continued
Author Peat (1991) [125]
N
Dose/Route
6 0.12 mg kg1 i.v.
Design
Outcome
Resting V_ E Resting PCO2; Resting rate; 5.5% CO2 challenge
#Resting V_ E ; #Resting rate; "Resting PCO2; # V_ E response to 5.5% CO2
10 0.1 mg kg1 i.v.
Resting rate; Resting PCO2,
#Resting rate; !Resting PCO2
Zhou (1993) [127]
16 0.15 mg kg1 i.v.
Resting V_ E ; Resting PCO2; CO2 rebreathe
#Resting V_ E ; "Resting PCO2; # V_ E vs CO2 slope
Lynn (1993) [128]
30 Washout following postoperative infusion
Resting PCO2; CO2 rebreathe
" Resting PCO2; # V_ E vs. CO2 slope
Morphine 6-glucuronide—no effect on resting variables; modest # V_ E response to CO2 challenge (less than half the effect of morphine) Slower onset and less pronounced effects than meperidine Effects more pronounced in Caucasian than in Chinese subjects; " morphine clearance in Chinese subjects Infants following heart surgery; depression correlates with serum concentration but not with age
Gross and Cerza
Hamunen (1993) [126]
Comment
Thompson (1995) [130] Borgbjerg (1996) [131] Dahan (1998) [132]
125 0.2 mg kg1 i.m.
10 0.14 mg kg1 10 0.2 mg kg1
CO2 rebreathe
# V_ E vs. CO2 slope
Resting PCO2; Resting V_ E CO2 rebreathe
"Resting PCO2; #Resting V_ E ! V_ E vs. CO2 slope; Right shift of V_ E vs. CO2 # V_ E vs. CO2 slope in women; ! V_ E vs. CO2 slope with right shift of V_ E vs. CO2 in men; Hypoxic response # in women and ! in men # V_ E response to CO2 Peripheral component and hypoxia of CO2 response and hypoxic response # more in women than men Peak effect at 3.5 h. Similar # V_ E response to acute hypoxia effect with subarachnoid morphine 0.3 mg suggests central effect Effect least in Caucasians, # V_ E vs. CO2 slope intermediate in Latinos, most pronounced in Native Indians; peak ventilatory depression at 3h
24 0.1 mg kg1 bolus; Steady-state CO2 30 mg kg1 h1 response; Dynamic end-tidal forcing of O2 infusion
Sarton (1999) [133]
18 0.1 mg kg1 bolus; Dynamic end-tidal 30 mg kg h1 forcing of PETCO2 infusion and PETO2
Bailey (2000) [134]
30 0.14 mg kg1 i.v.
Cepeda (2001) [135] 66 0.14 mg kg1 i.v.
Dynamic end-tidal forcing of PETO2 CO2 rebreathing
Chinese subjects more susceptible to depressant effects; V_ E response correlates with resting pulse Morphine 6-glucuronide—no effect on ventilation Effect counteracted by painful stimulation
Ventilatory Effects of Medications Used for Sedation
Houghton (1994) [129]
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increased to 0.4 mg kg1 or more, the slope of the CO2 response line begins to decrease significantly. Since at these higher doses morphine tends to produce sedation in addition to analgesia, it seems plausible that the observed decrease in slope is related to a decrease in the level of consciousness, similar to the effect of low doses of sedatives. In contrast, when the CO2 response is determined by rebreathing techniques, even low doses of morphine consistently appear to decrease the slope of the CO2 response curve (Table 14.6). This may be an artifact of the measurement technique rather than an effect of morphine itself. Specifically, the slope of the CO2 response as determined by the Read rebreathing method might not accurately reflect the slope as determined by steadystate methods. This reflects a changing gradient between arterial and chemoreceptor CO2 tensions related to changing cerebral blood flow during rebreathing measurements [138]. Additionally, the slope of the rebreathing CO2 response curve is critically dependent upon the difference between the resting PCO2 and the initial CO2 of the rebreathing system. If resting PCO2 is increased, but the same (e.g., 7%) CO2 concentration is initially used in the rebreathing system, the slope of the measured CO2 response curve tends to be decreased, even if the sensitivity of the chemoreceptor is unchanged [124]. Thus, studies in which the CO2 response was determined by rebreathing cannot distinguish between changes in the sensitivity of the chemoreceptor (i.e., the slope of the CO2 response curve) and changes in the set-point (i.e., parallel shifts in the curve without changes in slope). Even when administered intravenously, the onset of morphine-induced ventilatory depression is not immediate. For example, Hamunen observed that peak resting PCO2 occurred 12 min after a single dose of morphine 0.1 mg kg1 i.v. [126]. Using a rebreathing technique, Camporesi et al. determined that ventilatory depression peaks between 30 and 60 min following the same dose of morphine [121]. This helps to explain why modest doses of morphine, given alone, seldom cause respiratory arrest: as ventilatory drive becomes gradually depressed, there is sufficient time for PCO2 to increases and stimulate central ventilatory centers before hypoxemia develops. Interestingly, Camporesi et al. found that a single 0.1 mg kg1 dose of morphine may depress ventilatory drive for 10 h or more [121]. Morphine also significantly depresses hypoxic ventilatory response. Weil et al. found that morphine 0.1 mg kg1 s.c. significantly depressed the shape parameter A of the hypoxic ventilatory response, with peak depression occurring 60 min after injection [117]. More recently, Bailey et al. [134] found a significant depression in the acute response to a stepdecrease in oxygenation following i.v. morphine 0.14 mg kg1. The peak effect occurred at a median time of 3.5 h after injection, with complete recovery by 12 h after injection.
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Published studies do not support the conventional wisdom that patients at extremes of age have an increased susceptibility to the respiratory depressant effects of morphine. Arunasalam et al. found equivalent decreases in resting ventilation and increases in PCO2 20 min after morphine 0.15 mg kg1 i.v. in young (28–37 yr) and old (65–82 yr) patients [122]. However, the elderly patients appeared more likely to develop partial airway obstruction after morphine. Daykin et al. found that morphine 0.15 mg kg1 i.v. decreased the slope of the V_ E response to CO2 by 21% in young patients (18–29 yr) and 33% in elderly patients (66–85 yr) [123]. While this suggests a slightly increased susceptibility of elderly patients to morphine-induced ventilatory depression, the difference was not statistically significant. Lynn et al. correlated CO2 response slopes with morphine serum levels in pediatric patients ranging in age from 2 to 570 days [128]. They found no age-related differences in the respiratory depressant effect of morphine; however, they did observe that even at relatively low plasma concentrations (515 ng ml1), some infants had essentially no ventilatory response to CO2. Dahan et al. found that morphine affects the ventilatory response to CO2 differently in men compared with women [132]. They found that in men, a morphine infusion (0.1 mg kg1 load followed by 30 mg kg1 h1 infusion) had no effect on the slope of the CO2 response; there was, however, a 4 mm Hg rightward shift of the CO2 response. In contrast, the slope of the CO2 response decreased by 28% in women, despite an equivalent clinical level of sedation. Hypoxic sensitivity was reduced by 50% in women, but remained unaffected in men, suggesting the peripheral chemoreceptors of women are more susceptible to opioid-induced depression. In a followup study, Sarton et al. used dynamic end-tidal forcing to dissect the CO2 ventilatory response into its rapidly and slowly responding components, corresponding to peripheral and central chemoreceptor control, respectively [133]. They found that in men, a morphine infusion (0.1 mg kg1 load followed by 30 mg kg1 h1 infusion) depressed the central but not the peripheral component of the CO2 ventilatory response, while in women both the central and peripheral components were significantly depressed, again suggesting increased susceptibility of the peripheral chemoreceptors in women. It has not been established whether these findings translate to an increased susceptibility of women to develop respiratory compromise at any given level of moderate or deep sedation. Ethnicity may also affect the ventilatory effects of morphine. In a study performed in Colombia, Cepeda et al. found that morphine 0.14 mg kg1 decreased the slope of the CO2 response by 16% in Caucasian men, 24% in men of Hispanic extraction, and 40% in native ‘Indian’ men [135]. These differences were unrelated to differences in morphine metabolism or in the concentrations of the active metabolites morphine3-glucuronide and morphine-6-glucuronide. Zhou et al. found that
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morphine 0.15 mg kg1 depressed the slope of the CO2 ventilatory response more in Caucasian than in Chinese men [127]. This finding cannot be entirely explained by differences in morphine metabolism between the two groups, since at any given plasma concentration of morphine, ventilatory depression was greater in the Caucasian men. In contrast, Houghton et al. found that Chinese men were more susceptible to the ventilatory depressant effects of morphine than Caucasian men [129]. However, when individuals with high baseline heart rates were excluded, the ethnic differences disappeared. The authors suggest that the ethnic differences may have been related to different levels of physical fitness between the groups. These data support the clinical impression that Hispanic, Native American (and possibly Afro-American) patients may, under some circumstances be more susceptible than Caucasians to the ventilatory depressant effects of opioids. B. Meperidine
Meperidine is commonly used to provide analgesia in patients requiring moderate or deep sedation, particularly for endoscopic procedures; its potency is approximately one-tenth that of morphine. In doses typically used for moderate and deep sedation, its hemodynamic and respiratory effects are similar to those of morphine. However, in larger doses (41 mg kg1) meperidine may be associated with an increased incidence of histamine release with associated tachycardia and hypotension [147]. In addition, accumulation of the metabolite normeperidine may be associated with central nervous system activation and seizure activity, particularly in patients with renal insufficiency [148]. Studies delineating the ventilatory effects of meperidine are summarized in Table 14.7. Early studies by Campbell et al. demonstrated decreased ventilation in response to a CO2 challenge following i.v. meperidine [116,139]. While the time course of the effect was not evaluated, they found that the respiratory depressant effect of meperidine 61 mg was equivalent to that produced by morphine 10 mg [116]. Downes et al. first evaluated the onset and extent of meperidine-induced ventilatory depression; they continuously monitored ventilatory depression using an isohypercapnic technique [140]. They found that following meperidine 1.05 mg kg1 i.m., a decrease in V_ E at PETCO2 ¼ 46 mm Hg could be detected within 10 min; the peak effect, a 53% decrease, occurred 90 min after injection, and respiratory depression persisted for at least 4 h after injection. The relatively slow onset and long duration of meperidine-induced ventilatory depression in this study may be related to the fact that meperidine was administered i.m. rather than i.v. In contrast, using a rebreathing technique, Paulson et al. demonstrated that meperidine 1.4 mg kg1 i.v. reduced the slope of the V_ E response to
Studies Evaluating the Effects of Meperidine on Ventilatory Drive
Author
N
Dose/Route 0.7–2.1 mg kg1 i.v. 0.7–1.7 mg kg1 i.v. 1.05 mg kg1 i.m. 1.65 mg kg1 i.m.
Design Single CO2 challenge Single CO2 challenge Isohypercapnic CO2 rebreathe
Campbell (1964) [116] Campbell (1965) [139] Downes (1967) [140] Rouge (1969) [141]
6 4 6 5
Reier (1970) [142]
6 0.7–1.4 mg kg1 i.m.
CO2 rebreathe
Kaufman (1979) [143]
5 0.55–1.10 mg kg1 i.v.
CO2 rebreathe; isohypercarbia CO2 rebreathe
Rigg (1981) [144]
10 Target controlled 100–1200 ng ml1 i.v.
Paulson (1983) [145] Hersh (1985) [146]
20 1.4 mg kg1 i.v. 8 0.35–0.7 mg kg1 i.v.
Hamunen (1993) [126] 10 0.67 mg kg1
Outcome "PaCO2 # V_ E ; !PaCO2 # V_ E ! V_ E vs. CO2 slope; Right shift of V_ E vs. CO2 ! V_ E vs. CO2 slope; Right shift of V_ E vs. CO2 # V_ E vs. CO2 slope # V_ E vs. CO2 slope
CO2 rebreathe Steady-state CO2 response
# V_ E vs. CO2 slope Right shift of V_ E vs. CO2
Resting rate; Resting PCO2
#Resting rate; "Resting PCO2
Comment
Maximum effect at 90 min
Peak effect at 8 min Parallel shift at lower doses; # slope at higher doses (dose-dependent) Peak effect at 60 min Most pronounced effect ‘immediately post drug’; dose-dependent effect Peak CO2 at 6 min
Ventilatory Effects of Medications Used for Sedation
Table 14.7
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Gross and Cerza
CO2 by 40%; the peak effect occurred within 15 min, and the depression persisted for at least 2 h [145]. Similarly, Hersh et al. found that i.v. meperidine 0.7 mg kg1 caused a right shift in the CO2 response curve as determined by steady-state methodology; the peak effect occurred 30 min after the drug was injected, and respiratory depression persisted for at least 60 min [146]. Most recently, Hamunen et al. compared the onset and duration of ventilatory depression following i.v. morphine and meperidine in children; they found that respiratory depression, as assessed by resting PETCO2 and oxygen saturation, was more pronounced and peaked more rapidly following meperidine 0.6 mg kg1 than following morphine 0.1 mg kg1 [126]. In summary, the ventilatory effects of meperidine are similar to those of an equipotent dose of morphine. Since the usual doses are relatively low, the primary effect appears to be displacement of the CO2 response curve rather than a change in its slope. The onset of ventilatory depression may be more rapid with meperidine, as a result of its increased lipid solubility. Due to the potential for adverse hemodynamic and central nervous system effects, the effects of higher doses of meperidine (41.5 mg kg1) on ventilatory drive have not been evaluated. C. Fentanyl
Due to its high lipid solubility, fentanyl crosses the blood–brain barrier rapidly, accounting for the rapid onset of its analgesic and ventilatory depressant effects. It also redistributes rapidly into muscle and fat, so the central-nervous-system effects of a single dose of fentanyl are of shorter duration than those following morphine or meperidine. Fentanyl is approximately 100 times more potent than morphine and 1,000 times more potent than meperidine. The ventilatory effects of fentanyl are summarized in Table 14.8. The rapid onset and offset of fentanyl-induced ventilatory depression was clearly demonstrated in the study of Harper et al. [149]. These investigators found that even small doses of fentanyl (1.5 mg kg1) significantly increased resting PETCO2 within 2 min of i.v. injection. The peak effect occurred 5 min after injection, and within 2 h, PETCO2 had returned to baseline. Using a rebreathing technique, these investigators observed a 30% decrease in the slope of the ventilatory response at 5 min after fentanyl injection, with a return to baseline within an hour after injection. With larger fentanyl doses (up to 9 mg kg1), the onset of respiratory depression was equally rapid; however the slope of the ventilatory response decreased by 53%, and ventilatory drive remained depressed for at least 3 h. Using an isohypercapnic technique, Kaufman et al. [143] found that fentanyl 1.4 mg kg1 significantly decreased
Studies Evaluating the Effects of Fentanyl on Ventilatory Drive
Author
N
Dose/Route
Design
Downes (1967) [140] Harper (1976) [149] Kaufman (1979) [143]
6 10 5
1.4–2.8 mg kg1 i.m. 1.5–9 mg kg1 i.v. 0.7–1.4 mg kg1 i.v.
Isohypercapnic CO2 rebreathe CO2 rebreathe
Stoeckel (1982) [150]
7
8 mg kg1 i.v.
CO2 rebreathe
V_ E V_ E vs. CO2 slope V_ E vs. CO2 slope (high dose); Right shift V_ E vs. CO2 (low dose) # V_ E vs. CO2 slope
1.5–3 mg kg1 i.v.
CO2 rebreathe
Right shift V_ E vs. CO2
3 mg kg1 i.m.
CO2 rebreathe
! V_ E vs. CO2 slope
1–4 mg kg1 i.v.
CO2 rebreathe
# V_ E vs. CO2 slope
Scamman (1984) [151]
16
Negre (1987) [152]
9
Bailey (1990) [153]
30
Outcome # # #
Comment Peak effect at 30 min Peak effect at 5 min Peak effect at 8 min
Secondary respiratory depression (1 h post injection) in 4 subjects Peak at 4 min; recovery half-time 64 min Significant decrease in V_ E response with same dose epidurally Duration (4 h) significantly longer than for equivalent dose of sufentanil (30 min)
Ventilatory Effects of Medications Used for Sedation
Table 14.8
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Gross and Cerza
CO2-stimulated ventilation, with a peak effect at about 8 min; the slope of the CO2 response curve, as determined by CO2 rebreathing, decreased by 28%. A smaller dose of fentanyl, 0.7 mg kg1, did not significantly affect the slope of the CO2 response but did cause a right shift of the CO2 response curve. Bailey et al. found that fentanyl (1–4 mg kg1) significantly decreased the slope of both the ventilatory and the P0.1 responses to CO2 as determined by a rebreathing technique. The peak respiratory depression occurred within 5 min of drug injection [153]. In summary, the ventilatory effects of fentanyl parallel those of other m-opioid agonists. Low doses shift the CO2 response curve to the right, while higher doses decrease its slope; in anesthetic doses (30–50 mg kg1) fentanyl may cause chest-wall and laryngeal rigidity. The key distinction is that the onset and duration of fentanyl-induced ventilatory depression are both shorter than those of morphine or meperidine. Rapid-onset depression of ventilatory drive, particularly in sedated patients, may result in apnea as described below. D. Remifentanil
Remifentanil is an ultra-short-duration opioid, which is metabolized in situ by nonspecific plasma esterases. Using an isohypercapnic technique, Glass et al. found that a whole-blood remifentanil concentration of 1.2 ng ml1 produced a 50% decrease in V_ E while breathing 7.5% CO2 [154]. These investigators did not study the time-course of remifentanilinduced ventilatory depression. Babenco et al. studied the effect of a 0.5 mg kg1 bolus dose of remifentanil using a dual isohypercapnic technique [155]. They found a 72% decrease in the slope of the CO2 response curve 2 min after remifentanil injection; within 15 min, the slope returned to baseline (Figure 14.5). Apnea did not occur because of the stimulating effect of the artificially maintained elevated CO2 tension. These authors estimated that the EC50 for remifentanil-induced ventilatory depression is 1.1 ng ml1, and that the onset half-time (T1/2 ke0) was 2.9 min. Amin et al. studied the effect of remifentanil on the ventilatory response to hypoxia [156]. They found that with an infusion at the rate of 0.025 mg kg1 min1, remifentanil decreased the slope of the hypoxic ventilatory response (V_ E vs. SaO2) by 40%. With a remifentanil dose of 0.1 mg kg1 min1, hypoxic response decreased by 90%. In both cases, hypoxic ventilatory response returned to baseline within minutes after the remifentanil infusion was discontinued. Using a novel approach which accounted for time-related changes in both PCO2 and remifentanil-induced ventilatory depression, Bouillon et al. developed a model to predict the ventilatory effects of various
Ventilatory Effects of Medications Used for Sedation
551
20 0 18 15 16 10
14 · VE (l·min−1)
8 12 1 10
5 2
8 6
Time (min)
4 2 0 46
48
50
52
54
56
58
PETCO (mm Hg) 2
Figure 14.5 Constructed CO2 response curves by the dual isohypercapnic technique at the indicated times following injection of remifentanil 0.5 mg kg1 i.v (Data from Ref. 155).
remifentanil administration schemes on the time course of unstimulated V_ E and PCO2 [157]. This is analogous to the clinical situation where a drug such as remifentanil would be given to a patient whose breathing is not stimulated by exogenous CO2. This model predicts that a 1 mg kg1 bolus dose of remifentanil, which results in a peak blood level of 5 ng ml1, will cause an 80% decrease in alveolar ventilation; ventilation recovers as PCO2 rises and remifentanil concentrations fall. In contrast, an infusion designed to reach a similar blood concentration over 15 min causes a less dramatic initial decrease in ventilation, since the gradually increasing ventilatory depressant effect of the opioid is partially offset by the simultaneous increase in PCO2. Thus, to avoid hypoxemia resulting from hypoventilation, it is recommended that when remifentanil is used for moderate or deep sedation, it should be administered by continuous infusion rather than intermittent bolus doses. Furthermore, care must be taken to ensure that no remifentanil remains in the dead space of the intravenous tubing, where it might be accidentally administered if the infusion rate were subsequently increased [158].
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Gross and Cerza E.
Antagonism of Opioid-induced Ventilatory Depression
Naloxone
Naloxone is a pure m-opioid antagonist, which is devoid of respiratorydepressant effects. In fact, naloxone may stimulate ventilation both in normal volunteers and in patients with chronic obstructive pulmonary disease. In healthy volunteers, Akiyama et al. found a significant increase in hypoxic ventilatory response, as measured during modest hypercarbia, following naloxone 3 mg [159]. Similarly, Santiago et al. found that naloxone 2 mg i.v. more than doubled the ventilatory and P0.1 responses to hypoxia in patients with COPD [160]. These findings suggest that endogenous opioids such as beta endorphins may play a role in modulating resting hypoxic ventilatory drive. To most efficiently determine the effect of naloxone in reversing opioid-induced ventilatory depression, the naloxone can be given during a continuous infusion of an opioid agonist. In this way, the onset and duration of reversal can be assessed while avoiding the confounding effect of decreasing opioid-agonist concentrations. Glass et al. administered naloxone 1–8 mg kg1 to volunteers receiving a continuous fentanyl infusion (plasma concentration 2 ng ml1) [161]. They found a dose of 4 mg kg1 was required to completely antagonize fentanyl-induced depression of the ventilatory response to CO2. They also found that the duration of reversal was dose-dependent. With 4 mg kg1 of naloxone, reversal lasted approximately 70 min; with 8 mg kg1, reversal lasted almost twice as long. Amin et al. studied the effect of naloxone on morphine-induced depression of hypoxic ventilatory drive [162]. They found that morphine 0.125 mg kg1 decreased the slope of the hypoxic ventilatory response by 54%. Following naloxone 5 mg kg1, hypoxic ventilatory response returned to near its baseline value; however an hour later, the hypoxic ventilatory response decreased, indicating that the naloxone effect was waning more rapidly than that of the previously administered morphine. The relatively short duration of action of i.v. naloxone, when compared with the duration of ventilatory depression with morphine, meperidine or fentanyl, has led to the recommendation that patients who require reversal of an opioid with naloxone following moderate sedation be medically observed for at least one hour to ensure that respiratory depression does not recur [6]. Physostigmine
Snir-Mor et al. [163] used a rebreathing technique to determine that morphine 0.17 mg kg1 decreased the slope of the ventilatory response to
Ventilatory Effects of Medications Used for Sedation
553
CO2 by 37%; after physostigmine (13–33 mg kg1), the slope returned to its baseline value. In contrast, in the absence of morphine, these investigators found that physostigmine decreased the slope of the CO2 response. This, along with the subsequent finding of Bourke et al. that physostigmine did not antagonize morphine-induced depression of ventilation during isohypercapnia [28], suggest that physostigmine is probably not effective for this indication. Doxapram
Doxapram, a centrally acting respiratory stimulant, significantly increases the ventilatory response to CO2. Ramamurthy et al. demonstrated that following a dose of 2 mg kg1 i.m., doxapram shifted the CO2 response curve to the left by as much as 14 mm Hg, with the effect lasting at least 3 h [164]. As expected, these investigators also found that meperidine 1 mg kg1 caused an 8 mm Hg right shift of the CO2 response curve. When administered together, they found that doxapram completely prevented the meperidine-induced shift of the CO2 response curve. Similarly, Randall et al. found that oral doxapram 300–600 mg partially antagonized the ventilatory depressant effect of morphine 0.12 mg kg1 i.m. [165]. This interaction between doxapram and opioids probably represents the sum of two antagonistic ventilatory effects rather than true reversal of opioid-induced ventilatory depression.
IV.
Drug Combinations
A. Benzodiazepines Plus Opioids
A combination of a benzodiazepine and an opioid is frequently administered to provide moderate or deep sedation for patients undergoing invasive procedures such as endoscopy and angiography. The benzodiazepine provides sedation and amnesia, while the opioid provides analgesia and reduces the likelihood of disinhibition during painful stimulation. Early studies of the respiratory effects of such combinations revealed that the benzodiazepine component had negligible ventilatory effects when compared with those of the opioid. For example, Cohen et al. found that diazepam 0.133–0.266 mg kg1 had a negligible effect on the displacement of the V_ E response to CO2 as determined by a rebreathing technique, while meperidine 0.5 mg kg1 shifted the CO2 response curve to the right by 7 mm Hg. When meperidine was combined with the lower dose of diazepam, the shift in the CO2 response was slightly less than that observed with meperidine alone, leading the authors to speculate that the depressant effect of meperidine had been counteracted.
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However, when meperidine was combined with the larger dose of diazepam, a 20 mm Hg right shift was observed, suggesting potentiation of ventilatory depression and leading the authors to suggest ‘that caution be exercised in the intravenous administration of this drug combination’ [166]. The study of Lehmann et al. suggests a possible mechanism for such an interaction [107]. They found that following alfentanil 15 mg kg1 i.v., the CO2 response, as determined by rebreathing, was shifted to the right without change in slope. When the study was repeated following premedication with diazepam 5 mg i.v., the slope of the CO2 response decreased by 15%. Thus, the sedation associated with diazepam may potentiate opioid-induced ventilatory depression by causing a change in slope rather than just a parallel right-shift of the curve. (A change in slope has more significant consequences, since it portends a decreased ability to respond at high CO2 tensions.) Bailey et al. performed a systematic evaluation of the interaction between midazolam and fentanyl using both repeated CO2 rebreathing measurements and resting measurements of oxygen saturation during room air breathing [41]. They found that midazolam 0.05 mg kg1 i.v. did not significantly affect the CO2 response curve, and there were no instances of hypoxemia (SpO2 5 90%) during unstimulated breathing. In contrast, fentanyl 2 mg kg1 i.v. decreased the slope of the CO2 response by approximately 50%, and six of 12 subjects had one or more hypoxemic episodes. When the drugs were combined, the effect on the CO2 response was indistinguishable from that of fentanyl alone; however, in this case 11 of 12 subjects had one or more episodes of hypoxemia. Of note, almost all of the hypoxemic episodes occurred within 5 min of drug administration. The authors suggest that the increased incidence of hypoxemia was related to the significant depression of the hypoxic ventilatory response by both benzodiazepines and opioids. Due to the rapid onset of midazolam and fentanyl, arterial oxygen partial pressure drops to critically low levels before blood CO2 tensions can rise adequately to stimulate breathing [41]. Using a steady-state technique, Gross et al. found that even very low plasma concentrations of alfentanil (9 ng ml1) decreased the slope of the ventilatory response to CO2 from 2.14 to 1.43 l min1 mm Hg1 [50]. Midazolam, titrated to a state of moderate sedation, further depressed the slope to 0.87 l min1 mm Hg1, suggesting an additive or synergistic effect. Similarly, alfentanil decreased the slope of the hypoxic ventilatory response, as determined by isocapnic rebreathing (PETCO2 ¼ 46–48 mm Hg), from 0.73 to 0.51 l min1 %Sp1 O2 ; midazolam further decreased hypoxic drive to 0.26 l min1 %Sp1 . The depressant effect of alfentanil was related O2 to a decrease in both respiratory rate and tidal volume, while the effect of midazolam was on tidal volume alone.
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B. Fentanyl Plus Droperidol
Mulroy et al. administered 3 ml of InnovarÕ (fentanyl 50 mg ml1 þ droperidol 2.5 mg ml1) to patients undergoing spinal, epidural or brachial plexus anesthesia. They found that the slope of the CO2 response decreased by 15% within 30 min of drug administration. By 90 min, ventilatory drive returned to baseline [167]. Becker et al. administered fentanyl (50 mg ml1) or InnovarÕ 0.03 ml kg1 as an anesthetic premedication [168]. They found that the slope of the CO2 response decreased by 37% following fentanyl, and by 30% following InnovarÕ . They concluded that droperidol does not appear to augment fentanyl-induced ventilatory depression, and might partially counteract it. Harper et al. compared the effects of fentanyl (50 mg ml1) with those of an equal volume of InnovarÕ [149]. They found that fentanyl 3 mg kg1 decreased the slope of the ventilatory response to 52% of control within 5 min; 2 h later, it had returned to only 84%. When an equal volume of InnovarÕ was administered, the initial decrease in slope was the same; however, 2 h later the slope was 16% greater than baseline. This suggests that the ventilatory stimulating effect of droperidol (see above) persisted after the respiratory depressant effect of fentanyl had diminished. This is consistent with the relative clinical duration of the two drugs. C. Opioids Plus Tranquilizers
Tranquilizers such as hydroxyzine, promethazine, and prochlorperazine may be combined with an opioid to produce moderate or deep sedation; it was thought that such combinations could augment the opioid-induced analgesia without increasing the risk of respiratory depression. One such combination, known as DPT (DemerolÕ [meperidine], PhenerganÕ [promethazine], ThorazineÕ [chlorpromazine]), is sometimes used to provide sedation for children. Using a rebreathing technique, Gabathuler et al. demonstrated that hydroxyzine caused a significant right shift of the CO2 response curve which lasted at least 3 h, suggesting that addition of hydroxyzine to an opioid might worsen respiratory depression [169]. In contrast Zsigmond et al. found that hydroxyzine significantly increased resting PaO2, an indication of respiratory stimulation; however, when hydroxyzine was given with meperidine, they found that the depression of resting PaO2 was greater than that observed with meperidine alone [170]. The authors attributed this finding to the fact that most subjects fell asleep after receiving the combination of drugs. This apparent contradiction can be explained by the data of Olson et al. [171]. These investigators found that morphine significantly decreased the slope of the ventilatory response to CO2, but addition of prochlorperazine caused no further decrease in the CO2 response. Furthermore, they found that
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morphine significantly depressed the ventilatory response to isocapnic hypoxia, and prochlorperazine completely reversed this effect. Thus, it appears that the ventilatory effect of an opioid–tranquilizer combination may depend upon the conditions under which it is measured. As compared with an opioid alone, the CO2 response may be further blunted (perhaps as a result of decreased level of consciousness) while the hypoxic response may be augmented. Note that these results are similar to those observed with a combination of an opioid and diphenhydramine (see above) [111]. D. Ketamine Plus Benzodiazepine
When ketamine is used for moderate or deep sedation, a benzodiazepine is frequently coadministered to reduce the likelihood that any dysphoric reactions will be recalled. Zsigmond et al. found an additive respiratory depressant effect between ketamine and diazepam [172]. When administered alone, ketamine 2 mg kg1 i.v. decreased resting PaO2 from 84 to 58 mm Hg; a similar decrease in PaO2, from 74 to 46 mm Hg was observed when the same dose of ketamine was administered 5 min after i.v. diazepam 0.2 mg kg1. Peak depression of PaO2 occurred within 2 min, and the ketamine component of respiratory depression lasted about 10 min; however, diazepam-treated patients continued to have lower PaO2 at this time. Parker et al. [173] administered midazolam (up to 0.1 mg kg1) followed by ketamine (1–2 mg kg1) to pediatric patients undergoing invasive (lumbar puncture, bone marrow aspiration) and non-invasive (radiotherapy, imaging) studies. Only four of these patients developed transient hypoxemia (SpO2 85%), which was readily treated by supplemental oxygen; however, 24 of 74 patients undergoing lumbar puncture developed mild hypoxemia (SpO2 88–94%) during neck flexion, which resolved with extension of the neck or supplemental oxygen. Similarly, Marx et al. found that only 17% of children receiving a combination of ketamine 1.5 mg kg1 and midazolam 0.05 mg kg1 developed hypoxemia (SpO2 5 92%) during painful procedures compared with 78% of those who received meperidine 2 mg kg1 plus midazolam 0.1 mg kg1; nevertheless, patients who received ketamine experienced less distress during their procedures [174]. E.
Ketamine Plus Opioid
Bourke et al. found that following i.v. morphine, ketamine caused a modest dose-related enhancement of respiratory depression [175]. Morphine 0.2 mg kg1 decreased V_ E during isohypercapnia by 10 l min1, and the maximum 3 mg kg1 dose of ketamine caused a further 5 l min1 decrease. Similarly, 0.4 mg kg1 of morphine decreased V_ E by 15 l min1 with ketamine again causing an additional 5 ml min1 decrease. Neither the
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morphine nor the ketamine changed the slope of the CO2 response, consistent with the parallel shift in CO2 response observed by Hamza et al. [103]. In contrast, Mildh et al. used respiratory impedance plethysmography to determine that fentanyl 2 mg kg1 plus ketamine 0.25 mg kg1 caused a smaller decrease in resting minute ventilation than fentanyl 2 mg kg1 given alone [176]. The maximum decrease in oxygen saturation was 6% with both treatments; this discrepancy was probably related to an increase in oxygen consumption when ketamine was administered. These results most likely differ from those of Bourke et al. by virtue of the low dose of ketamine; in fact, Bourke found that ketamine 0.4 mg kg1 had no effect on ventilation [175]. F. Ketamine Plus Propofol
The rationale for using ketamine–propofol mixtures for moderate and deep sedation is that low doses of ketamine can provide analgesia without potentiating the respiratory depressant effects of propofol. After premedicating patients with midazolam 1–3 mg and fentanyl 50 mcg, Mortero et al. administered either propofol (10 mg ml1) or a mixture of propofol (9.8 mg ml1) and ketamine (0.98 mg ml1) to achieve a level of deep sedation [177]. Patients in the propofol and propofol–ketamine groups required similar propofol doses (38 vs. 33 mg kg1 min1), yet PETCO2 increased in patients receiving propofol alone while it decreased in those receiving the propofol–ketamine mixture. This difference was related to a higher respiratory rate in those receiving the propofol–ketamine mixture. In a similar study, Badrinath et al. found that patients who received propofol were more likely to develop airway obstruction than those who received a propofol–ketamine mixture for moderate sedation (52% vs. 20%) [178].
V.
Strategies for Minimizing Respiratory Risks of Sedation
A. Dose Titration
Conventional wisdom holds that titration of sedative and analgesic medications to appropriate end points based on patients’ level of consciousness, respiratory and cardiovascular functions, reduces the risk of hypoventilation, airway obstruction, hypoxemia, and hypotension. Thus, the ASA Guidelines for Sedation and Analgesia by non-Anesthesiologists recommend administering incremental doses of sedatives to relieve anxiety and opioids to alleviate procedural discomfort [5,6]. However, there are some data suggesting that dose titration alone may not be sufficient to eliminate the risks associated with moderate and deep sedation.
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Alexander et al. found that when compared with a bolus dose, administration of midazolam 0.1 mg kg1 over 5 min delayed the decrease in slope of the CO2 response as assessed by the dual-isohypercapnic technique [44]. However, within 3 min after the infusion was complete, identical degrees of respiratory depression were observed. These findings regarding the CO2 response may not reflect the effect on oxygenation in the clinical situation; slow administration would allow time for CO2 to increase, partially offsetting the ventilatory depressant effect of the midazolam infusion [157]. In a surprising study, Morrow et al. compared the ventilatory effects of fixed, bolus doses of meperidine and midazolam with those of clinically titrated doses of the same medications in patients undergoing colonoscopy [51]. They found that patients in the dose titration group were more likely than those in the bolus dose group to develop hypoxemia (65% vs. 31%) or to require supplemental oxygen (44% vs. 14%). Although patients were equally satisfied after either regimen, procedures took longer to complete when dose titration was used. These findings may be related to the fact that during dose titration, additional doses were administered at 3 min intervals while the peak effect of meperidine may not occur for 10 min or more. Thus, it is possible that titration resulted in a cumulative overdose and subsequent respiratory depression. This is consistent with the fact that patients in the dose titration group received a significantly larger total meperidine dose than those in the bolus dose group (83 vs. 67 mg). This emphasizes the need to wait until the peak effect of each dose of a sedative or analgesic medication occurs before administering an additional dose when titration is used. B. Supplemental Oxygen
Administration of supplemental oxygen during moderate or deep sedation reduces the risk of hypoxemia. Gross et al. [179] demonstrated that during midazolam–meperidine sedation for colonoscopy, the risk of hypoxemia (SpO2 5 90%) was significantly lower in patients receiving nasal O2 at 2 l min1 (36% vs. 79%). Furthermore, patients receiving supplemental O2 averaged less than 1 min of hypoxemia during 41 min of colonoscopy; in contrast, those breathing room air averaged 9.7 min of hypoxemia during 38 min of colonoscopy. Administration of supplemental oxygen will delay the recognition of hypoventilation or apnea by pulse oximetry. However, this is not a good reason to avoid the routine use of supplemental oxygen. Rather, it emphasizes the need to use an independent monitor of ventilatory function such as observation, auscultation, or capnography to detect respiratory insufficiency before hypoxemia ensues [180]. Used in this way, supplemental oxygen provides an increased margin of safety by allowing additional time
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between the onset of ventilatory insufficiency and the development of hypoxemia. VI.
Conclusion
All of the medications used to provide moderate or deep sedation have a potential to decrease respiratory drive and/or predispose patients to develop airway obstruction. While it may be possible to minimize the risk of adverse outcomes through a thorough understanding of the respiratory pharmacodynamics of the medications being used, the potential for lifethreatening respiratory compromise is always present. Hence, patients receiving procedural sedation must be continuously monitored to detect the onset of such adverse events before irreversible harm ensues. Furthermore, practitioners administering sedative and analgesic medications for procedural sedation must have the skills necessary to rescue patients whose level of sedation becomes deeper than intended, resulting in respiratory compromise. Finally, it is important to recognize that drug-induced respiratory depression may persist after the diagnostic or therapeutic procedure is complete, so patients must be monitored in a medical setting until the effects of sedative and analgesic medications have dissipated. References 1. 2. 3.
4. 5.
6.
7. 8.
Monheim, L.M., Analgesia in dentistry now and in the future, Anesth. Prog. 8, 100–103, 1971. Lee, B., The dental auxiliary and oxygen-nitrous conscious sedation, The Dent. Assistant 42, 15–17, 1973. Bennett, C.R., ed., Conscious sedation in dentistry, in Monheim’s Local Anesthesia and Pain Control in Dental Practice, 7th ed., Mosby, St. Louis, pp. 267–287, 1984. U.S. is Asked to Sharply Limit Use of Sedative, New York Times, Feb. 14, 1988, p. 37. Practice guidelines for sedation and analgesia by non-anesthesiologists, A report by the American Society of Anesthesiologists Task Force on Sedation and Analgesia by Non-Anesthesiologists, Anesthesiology 84, 459–471, 1996. Practice guidelines for sedation and analgesia by non-anesthesiologists, An Updated Report by the American Society of Anesthesiologists Task Force on Sedation and Analgesia by Non-Anesthesiologists, Anesthesiology 96, 1004–1017, 2002. ValiumÕ Package Insert, Roche Pharmaceuticals, Manati, PR, 2001. Delpierre, S., Jammes, Y., Grimaud, C., Dugue, P., Arnaud, A. and Charpin, J., Influence of anxiolytic drugs (prazepam and diazepam) on respiratory center output and CO2 chemosensitivity in patients with lung diseases, Respiration 42, 15–20, 1981.
560 9. 10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21. 22. 23.
24. 25.
Gross and Cerza Mak, K.H., Wang, Y.T. and Cheong, T.H., The effect of oral midazolam and diazepam on respiration in normal subjects, Eur. Respir. J. 6, 42–47, 1993. Gilmartin, J.J., Corris, P.A., Stone, T.N., Veale, D. and Gibson, G.J., Effects of diazepam and chlormethiazole on ventilatory control in normal subjects, Br. J. Clin. Pharmacol. 25, 766–770, 1988. Utting, H.J. and Pleuvry, B.J., Benzoctamine—a study of the respiratory effects of oral doses in human volunteers and interactions with morphine in mice, Br. J. Anaesth. 47, 987–992, 1975. Rapoport, D.M., Greenberg, H.E. and Goldring, R.M., Differing effects of the anxiolytic agents buspirone and diazepam on control of breathing, Clin. Pharmacol. Ther. 49, 394–401, 1991. Beaupre, A., Soucy, R., Phillips, R. and Bourgouin, J., Respiratory center output following zopiclone or diazepam administration in patients with pulmonary disease, Respiration 54, 235–240, 1988. Leiter, J.C., Knuth, S.L., Krol, R.C. and Bartlett, D., Jr., The effect of diazepam on genioglossal muscle activity in normal human subjects, Am. Rev. Respir. Dis. 132, 216–219, 1985. Philip-Joe¨t, F., Marc, I. and Se´rie`s, F., Effects of genioglossus response to negative airway pressure on upper airway collapsibility during sleep, J. Appl. Physiol. 80, 1466–1474, 1996. Steen, S.N., Weitzner, S.W., Amaha, K. and Martinez, L.R., The effect of diazepam on the respiratory response to carbon dioxide, Can. Anaesth. Soc. J. 13, 374–377, 1966. Dalen, J.E., Evans, G.L., Banas, J.S., Jr., Brooks, H.S., Paraskos, J.A. and Dexter, L., The hemodynamic and respiratory effects of diazepam (ValiumÕ ), Anesthesiology 30, 259–263, 1969. Catchlove, R.F.H. and Kafer, E.R., The effects of diazepam on the ventilatory response to carbon dioxide and on steady-state gas exchange, Anesthesiology 34, 9–13, 1971. Catchlove, R.F.H. and Kafer, E.R., The effects of diazepam on respiration in patients with obstructive pulmonary disease, Anesthesiology 34, 14–18, 1971. Cegla, U.H., The use of CO2 response curves to determine the respirationdepressant action of drugs with diazepam as an example, Pneumonologie 149, 219–228, 1973. Gasser, J.C. and Bellville, J.W., The respiratory effects of hydroxyzine, diazepam and pentazocine in man, Anaesthesia 31, 718–723, 1976. Lakshminarayan, S., Sahn, S.A., Hudson, L.D. and Weil, J.V., Effect of diazepam on ventilatory responses, Clin. Pharm. Ther. 20, 178–183, 1976. Jordan, C., Tech, B., Lehane, J.R. and Jones, J.G., Respiratory depression following diazepam: Reversal with high-dose naloxone, Anesthesiology 53, 293– 298, 1980. Forster, A., Gardaz, J., Suter, P.M. and Gamperle, M., Respiratory depression by midazolam and diazepam, Anesthesiology 53, 494–497, 1980. Clergue, F., Desmonts, J.M., Duvaldestin, P., Delavault, E. and Saumon, G., Depression of respiratory drive by diazepam as premedication, Br. J. Anaesth. 53, 1059–1063, 1981.
Ventilatory Effects of Medications Used for Sedation 26. 27. 28.
29.
30.
31.
32.
33. 34.
35. 36.
37.
38.
39.
40. 41.
42.
561
Gross, J.B., Smith, L. and Smith, T.C., Time course of ventilatory response to carbon dioxide after intravenous diazepam, Anesthesiology 57, 18–21, 1982. Power, S.J., Morgan, M. and Chakrabarti, M.K., Carbon dioxide response curves following midazolam and diazepam, Br. J. Anaesth. 55, 837–841, 1983. Bourke, D.L., Rosenberg, M. and Allen, P.D., Physostigmine: Effectiveness as an antagonist of respiratory depression and psychomotor effects caused by morphine or diazepam, Anesthesiology 61, 523–528, 1984. Spaulding, B.C., Choi, S.D., Gross, J.B., Apfelbaum, J.L. and Broderson, H., The effect of physostigmine on diazepam-induced ventilatory depression: A double-blind study, Anesthesiology 61, 551–554, 1984. Bailey, P.L., Andriano, K.P., Goldman, M., Stanley, T.H. and Pace, N.L., Variability of the respiratory response to diazepam, Anesthesiology 64, 560–565, 1986. Berggren, L., Eriksson, I. and Mollenholt, P., Changes in breathing pattern and chest wall mechanics after benzodiazepines in combination with meperidine, Acta Anesthesiol. Scand. 31, 381–386, 1987. Sunzel, M., Paalzow, L., Berggren, L. and Eriksson, I., Respiratory effects in relation to plasma levels of midazolam and diazepam, Br. J. Clin. Pharmacol. 25, 561–569, 1988. Mora, C.T., Torjman, M. and White, P.F., Effects of diazepam and flumazenil on sedation and hypoxic ventilatory response, Anesth. Analg. 68, 473–478, 1989. Zakko, S.F., Seifert, H.A. and Gross, J.B., A comparison of midazolam and diazepam for conscious sedation during colonoscopy in a prospective doubleblind study, Gastrointest. Endosc. 49, 684–689, 1999. Gross, J.B., Resting ventilation measurements may be misleading, Anesthesiology 61, 110, 1984. Bouillon, T., Schmidt, C., Garstka, G., Heimbach, D., Stafforst, D., Schwilden, H. and Hoeft, A., Pharmacokinetic–pharmacodynamic modeling of the respiratory depressant effect of alfentanil, Anesthesiology 91, 144–155, 1999. Gross, J.B. and Zebrowski, M.E., Carel, W.D., Gardner, S. and Smith, T.C., Time course of ventilatory depression after thiopental and midazolam in normal subjects and in patients with chronic obstructive pulmonary disease, Anesthesiology 58, 540–544, 1983. Forster, A., Morel, D., Bachmann, M. and Gemperle, M., Respiratory depressant effects of different doses of midazolam and lack of reversal with naloxone—a double-blind randomized study, Anesth. Analg. 62, 920–924, 1983. Power, S.J., Chakrabarti, M.K. and Whitman, J.G., Response to carbon dioxide after oral midazolam and pentobarbitone, Anaesthesia 39, 1183–1187, 1984. Alexander, C.M. and Gross, J.B., Sedative doses of midazolam depress hypoxic ventilatory responses in humans, Anesth. Analg. 67, 377–382, 1988. Bailey, P.L., Pace, N.L., Ashburn, M.A., Moll, J.W.B., East, K.A. and Stanley, T.H., Frequent hypoxemia and apnea after sedation with midazolam and fentanyl, Anesthesiology 73, 826–830, 1990. Dahan, A. and Ward, D.S., Effect of i.v. midazolam on the ventilatory response to sustained hypoxia in man, Br. J. Anaesth. 66, 454–457, 1991.
562 43. 44. 45.
46.
47.
48.
49.
50.
51.
52. 53.
54.
55.
56.
57.
Gross and Cerza Gross, J.B., Weller, R.S. and Conard, P., Flumazenil antagonism of midazolam-induced ventilatory depression, Anesthesiology 75, 179–185, 1991. Alexander, C.M. and Gross, J.B., Slow injection does not prevent midazolaminduced ventilatory depression, Anesth. Analg. 74, 260–264, 1992. Blouin, R.T., Conard, P.F., Perrault, S. and Gross, J.B., The effect of flumazenil on midazolam-induced depression of the ventilatory response to hypoxia during isohypercarbia, Anesthesiology 78, 635–641, 1993. Flo¨gel, C.M., Ward, D.S., Wada, D.R. and Ritter, J.W., The effects of largedose flumazenil on midazolam-induced ventilatory depression, Anesth. Analg. 77, 1207–1214, 1993. Nagyova, B., Dorrington, K.L. and Robbins, P.A., Effects of midazolam and flumazenil on ventilation during sustained hypoxia in humans, Respir. Physiol. 94, 51–59, 1993. Mora, C.T., Torjman, M. and White, P.F., Sedative and ventilatory effects of midazolam infusion: Effect of flumazenil reversal, Can. J. Anaesth. 42, 677–684, 1995. Litman, R.S., Kottra, J.A., Berkowitz, R.J. and Ward, D.S., Breathing patterns and levels of consciousness in children during administration of nitrous oxide after oral midazolam premedication, J. Oral Maxillofac. Surg. 55, 1372–1377, 1997. Gross, J.B., Blouin, R.T., Zandsberg, S., Conard, P.F. and Haussler, J., Effect of flumazenil on ventilatory drive during sedation with midazolam and alfentanil, Anesthesiology 85, 713–720, 1996. Morrow, J.B., Zuccaro, G., Conwell, D.L., Vargo, J.J., Dumont, J.A., Karafa, M. and Shay, S.S., Sedation for colonoscopy using a single bolus is safe, effective, and efficient: A prospective, randomized, double-blind study, Am. J. Gastroenterol. 95, 2242–2247, 2000. Montavers, P., Dureuil, B. and Desmonts, J.M., Effects of i.v. midazolam on upper airway resistance, Br. J. Anaesth. 68, 27–31, 1992. Montavers, P., Dureuil, B., Molliex, S. and Desmonts, J.M., Effects of intravenous midazolam on the work of breathing, Anesth. Analg. 79, 558–562, 1994. Kawauchi, Y., Oshima, T., Suzuki, S., Saitoh, Y. and Toyooka, H., Advancement of the mandible facilitates nasal breathing in human subjects sedated with midazolam, Can. J. Anaesth. 47, 215–219, 2000. Nozaki-Taguchi, N., Isono, S., Nishino, T., Numai, T. and Taguchi, N., Upper airway obstruction during midazolam sedation: Modification by nasal CPAP, Can. J. Anaesth. 42, 685–690, 1995. Oshima, T., Masaki, Y. and Toyooka, H., Flumazenil antagonizes midazolaminduced airway narrowing during nasal breathing in humans, Br. J. Anaesth. 82, 698–702, 1999. Carter, A.S., Bell, G.D., Coady, T., Lee, J. and Morden, A., Speed of reversal of midazolam-induced respiratory depression by flumazenil—a study in patients undergoing upper G.I. endoscopy, Acta Anesthesiol. Scand. 92, 59–64, 1990.
Ventilatory Effects of Medications Used for Sedation 58.
59. 60. 61. 62.
63.
64. 65.
66. 67.
68.
69.
70.
71.
72.
73.
563
Forster, A., Crettenand, G., Klopfenstein, C.E. and Morel, D.R., Absence of agonist effects of high-dose flumazenil on ventilation and psychometric performance in human volunteers, Anesth. Analg. 77, 980–984, 1993. Caldwell, C.B. and Gross, J.B., Physostigmine reversal of midazolam-induced sedation, Anesthesiology 57, 125–127, 1982. Larson, G.F., Hurlbert, B.J. and Wingard, D.W., Physostigmine reversal of diazepam-induced depression, Anesth. Analg. 56, 348–351, 1977. Gesell, R. and Hansen, E.T., Eserine, acetylcholine, atropine and nervous integration, Am. J. Physiol. 139, 371–385, 1943. Bloomfield, E.L., Masaryk, T.J., Caplin, A., Obuchoski, N.A., Schubert, A., Hayden, J., Ebrahim, Z.Y., Ruggieri, P.M., Goske, M.J. and Ross, J.S., Intravenous sedation for MR imaging of the brain and spine in children: Pentobarbital versus propofol, Radiology 5, 310–314, 1993. Finn, H., Cohen, R. and Steen, S.N., Comparison of the effects of RO 5-6901, pentobarbital, and meperidine on the respiratory response to carbon dioxide, Anesth. Analg. 49, 279–299, 1970. Murray, A., Bellville, J.W., Comer, W. and Danielson, L., Respiratory effects of quazepam and pentobarbital, J. Clin. Pharmacol. 27, 310–313, 1987. Brown, C.R., Forrest, W.H., Jr and Hayden, J., The respiratory effects of pentobarbital and secobarbital in clinical doses, J. Clin. Pharmacol. New Drugs 13, 28–35, 1973. Gasser, J.C., Kaufman, R.D. and Bellville, J.W., Respiratory effects of lorazepam, pentobarbital, and pentazocine, Clin. Pharmacol. Ther. 18, 170–174, 1975. Hirshman, C.A., McCollough, R.E., Cohen, P.J. and Weil, J.V., Effect of pentobarbitone on hypoxic ventilatory drive in man: Preliminary study, Br. J. Anaesth. 47, 963–968, 1975. Greenblatt, D.J., Allen, M.D., Harmatz, J.S., Noel, B.J. and Shader, R.I., Overdosage with pentobarbital and secobarbital: Assessment of factors related to outcome, J. Clin. Pharmacol. 19, 758–768, 1979. Audenaert, S.M., Wagner, Y., Montgomery, C.L., Lock, R.L., Colclough, G., Kuhn, R.J., Johnson, G.L. and Pedigo, N.W., Cardiorespiratory effects of premedication for children, Anesth. Analg. 80, 506–510, 1995. Larsson, L.E., Nilsson, K., Andreasson, S. and Ekstro¨m-Jodal, B., Effects of rectal thiopentone and methohexitone on carbon dioxide tension in infant anaesthesia with spontaneous ventilation, Acta Anaesthesiol. Scand. 31, 227–230, 1987. Daniels, A.L., Cote´, C.J. and Polaner, D.M., Continuous oxygen saturation monitoring following rectal methohexitone induction in paediatric patients, Can. J. Anaesth. 39, 27–30, 1992. Audenaert, S.M., Montgomery, C.L., Thompson, D.E. and Sutherland, J., A prospective study of rectal methohexital: Efficacy and side effects in 648 cases, Anesth. Analg. 81, 957–961, 1995. Kay, B., The measurement of occlusion pressure during anaesthesia. A comparison of the depression of respiratory drive by methohexitone and etomidate, Anaesthesia 34, 543–548, 1979.
564 74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
87.
88. 89.
Gross and Cerza Choi, S.D., Spaulding, B.C., Gross, J.B. and Apfelbaum, J.L., Comparison of the ventilatory effects of etomidate and methohexital, Anesthesiology 62, 442–447, 1985. Bickler, P.E., Dueck, R. and Prutow, R.J., Effects of barbiturate anesthesia on functional residual capacity and ribcage/diaphragm contributions to ventilation, Anesthesiology 68, 147–152, 1987. Wise, C.C., Robinson, J.S., Heath, M.J. and Tomlin, P.J., Physiological responses to intermittent methohexitone for conservative dentistry, Br. Med. J. 2, 540–543, 1969. Mackenzie, N. and Grant, I.S., Comparison of propofol with methohexitone in the provision of anaesthesia for surgery under regional blockade, Br. J. Anaesth. 57, 1167–1172, 1985. Allen, G.D., Kennedy, W.F., Jr., Everett, G. and Tolas, A.G., A comparison of the respiratory effects of methohexital and thiopental supplementation for outpatient dental anesthesia, Anesth. Analg. 48, 730–735, 1969. Duffin, J., Triscott, A. and Whitwam, J.G., The effect of halothane and thiopentone on ventilatory responses mediated by the peripheral chemoreceptors in man, Br. J. Anaesth. 48, 975–981, 1976. Rigg, J.R.A. and Goldsmith, C.H., Recovery of ventilatory response to carbon dioxide after thiopentone, morphine and fentanyl in man, Can. Anaesth. Soc. J. 23, 370–382, 1976. Knill, R.L., Bright, S. and Manninen, P., Hypoxic ventilatory responses during thiopentone sedation and anaesthesia in man, Can. Anaesth. Soc. J. 25, 366–372, 1978. Germain, M., Wahba, W.M. and Gillies, D.M.M., Ventilation following induction of general anesthesia by thiopentone, Can. Anaesth. Soc. J. 29, 100– 104, 1982. Taylor, M.B., Grounds, R.M., Mulroney, P.D. and Morgan, M., Ventilatory effects of propofol during induction of anaesthesia, Anaesthesia 41, 816–820, 1986. Grounds, R.M., Maxwell, D.L., Taylor, M.B., Aber, V. and Royston, D., Acute ventilatory changes during i.v. induction of anaesthesia with thiopentone or propofol in man, Br. J. Anaesth. 59, 1098–1102, 1987. Valtonen, M., Anaesthesia for computerized tomography of the brain in children: A comparison of propofol and thiopentone, Acta Anaesthesiol. Scand. 33, 170–173, 1989. Blouin, R.T., Conard, P.F. and Gross, J.B., Time course of ventilatory depression following induction doses of propofol and thiopental, Anesthesiology 75, 940–944, 1991. Spens, H.J. and Drummond, G.B., Ventilatory effects of eltanolone during induction of anaesthesia: Comparison with propofol and thiopentone, Br. J. Anaesth. 77, 194–199, 1996. Drummond, G.B., Influence of thiopentone on upper airway muscles, Br. J. Anaesth. 63, 12–21, 1989. Goodman, N.W., Black, A.M.S. and Carter, J.A., Some ventilatory effects of propofol as sole anaesthetic agent, Br. J. Anaesth. 59, 1497–1503, 1987.
Ventilatory Effects of Medications Used for Sedation 90.
91.
92.
93.
94.
95.
96. 97.
98.
99. 100.
101.
102. 103. 104.
105.
565
Allsop, P., Taylor, M.B., Grounds, R.M. and Morgan, M., Ventilatory effects of a propofol infusion using a method to rapidly achieve steady-state equilibrium, Eur. J. Anaesthesiol. 5, 293–303, 1988. Vangerven, M., Van Hemelrijck, J., Wouters, P., Vandermeersch, E. and Van Aken, H., Light anaesthesia with propofol for paediatric MRI, Anaesthesia 47, 706–707, 1992. Blouin, R.T., Seifert, H.A., Babenco, H.D., Conard, P.F. and Gross, J.B., Propofol depresses the hypoxic ventilatory response during conscious sedation and isohypercapnia, Anesthesiology 79, 1177–1182, 1993. Nagyova, B., Dorrington, K.L., Gill, E.W. and Robbins, P.A., Comparison of the effects of sub-hypnotic concentrations of propofol and halothane on the acute ventilatory response to hypoxia, Br. J. Anaesth. 75, 713–718, 1995. Nieuwenhuijs, D., Sarton, E., Teppema, L. and Dahan, A., Propofol for monitored anesthesia care: Implication on hypoxic control of cardiorespiratory responses, Anesthesiology 92, 46–54, 2000. Nieuwenhuijs, D., Sarton, E., Teppema, L.J., Krut, E., Olievier, I., van Kleef, J. and Dahan, A., Respiratory sites of action of propofol: Absence of depression of peripheral chemoreflex loop by low-dose propofol, Anesthesiology 95, 889–895, 2001. Goodman, N.W. and Black, A.M.S., Ventilatory effects of propofol infusion (letter), Eur. J. Anaesthesiol. 6, 397–398, 1989. Litman, R.S., Weissend, E.E., Shirer, D.A. and Ward, D.S., Morphologic changes in the upper airway of children during awakening from propofol administration, Anesthesiology 96, 607–611, 2002. Hammer, J., Reber, A., Trachsel, D. and Frei, F.J., Effect of jaw-thrust and continuous positive airway on tidal breathing in deeply sedated infants, J. Pediatr. 138, 826–830, 2001. McKeating, K., Bali, I.M. and Dundee, J.W., The effects of thiopentone and propofol on upper airway integrity, Anaesthesia 43, 638–640, 1988. Vargo, J.J., Zuccaro, G., Jr., Dumot, J.A., Shay, S.S., Conwell, D.L. and Morrow, J.B., Gastroenterologist-administered propofol for therapeutic upper endoscopy with graphic assessment of respiratory activity: A case series, Gastrointest. Endosc. 52, 250–255, 2000. Morel, D.R., Forster, A. and Gemperle, M., Noninvasive evaluation of breathing pattern and thoraco-abdominal motion following the infusion of ketamine or droperidol in humans, Anesthesiology 65, 392–398, 1986. Bourke, D.L., Malit, L.A. and Smith, T.C., Respiratory interactions of ketamine and morphine, Anesthesiology 66, 153–166, 1987. Hamza, J., Ecoffey, C. and Gross, J.B., Ventilatory response to CO2 following intravenous ketamine in children, Anesthesiology 70, 422–425, 1989. Mitchell, R.K., Koury, S.I. and Stone, C.K., Respiratory arrest after intramuscular ketamine in a 2-year-old child, Am. J. Emerg. Med. 14, 580–581, 1996. Smith, J.A. and Santer, L.J., Respiratory arrest following intramuscular ketamine injection in a 4-year-old child, Ann. Emerg. Med. 22, 613–615, 1993.
566
Gross and Cerza
106. Soroker, D., Barzilay, E., Konichezky, S. and Bruderman, I., Respiratory function following premedication with droperidol or diazepam, Anesth. Analg. 57, 695–699, 1978. 107. Lehmann, K.A. and Mainka, F., Ventilatory CO2-response after alfentanil and sedative premedication (etomidate, diazepam and droperidol). A comparative study with human volunteers, Acta Anaesth. Belgica 37, 3–13, 1986. 108. Prokocimer, P., Delavault, E., Rey, F., Lefevre, P., Mazze, R.I. and Desmonts, J.M., Effects of droperidol on respiratory drive in humans, Anesthesiology 59, 113–116, 1983. 109. Ward, D.S., Stimulation of hypoxic ventilatory drive by droperidol, Anesth. Analg. 63, 106–110, 1984. 110. Alexander, C.M., Seifert, H.A., Blouin, R.T., Conard, P.F. and Gross, J.B., Diphenhydramine enhances the interaction of hypercapnic and hypoxic ventilatory drive, Anesthesiology 80, 789–795, 1994. 111. Babenco, H.D., Blouin, R.T., Conard, P.F. and Gross, J.B., Diphenhydramine increases ventilatory drive during alfentanil infusion, Anesthesiology 89, 642–647, 1998. 112. Lees, M.H., Olsen, G.D., McGillard, K.L. and Newcomb, J.D., Sunderland, C.O., Chloral hydrate and the carbon dioxide chemoreceptor response: A study of puppies and infants, Pediatrics 70, 447–450, 1982. 113. Litman, R.S., Kottra, J.A., Verga, K.A., Berkowitz, R.J. and Ward, D.S., Chloral hydrate sedation: The additive sedative and respiratory depressant effects of nitrous oxide, Anesth. Analg. 86, 724–728, 1998. 114. Cote´, C.J., Karl, H.W., Notterman, D.A., Weinberg, J.A. and McCloskey, C., Adverse sedation events in pediatrics: Analysis of medications used for sedation, Pediatrics 106, 633–644, 2000. 115. Hershenson, M., Brouillette, R.T., Olsen, E. and Hunt, C.E., The effect of chloral hydrate on genioglossus and diaphragmatic activity, Pediatr. Res. 18, 516–519, 1984. 116. Campbell, D., Lister, R.E. and McNicol, G.W., Drug-induced respiratory depression in man, Clin. Pharmacol. Ther. 5, 193–200, 1964. 117. Weil, J.V., McCullough, R.E., Kline, J.S. and Sodal, I.E., Diminished ventilatory response to hypoxia and hypercapnia after morphine in normal man, N. Engl. J. Med. 292, 1103–1106, 1975. 118. Knill, R., Cosgrove, J.F., Olley, P.M. and Levison, H., Components of respiratory depression after narcotic premedication in adolescents, Can. Anaesth. Soc. J. 23, 449–458, 1976. 119. Rigg, J.R.A., Ventilatory effects and plasma concentration of morphine in man, Br. J. Anaesth. 50, 759–765, 1978. 120. Gal, T.J., DiFazio, C.A. and Moscicki, J., Analgesic and respiratory depressant activity of nalbuphine: A comparison with morphine, Anesthesiology 57, 367–374, 1982. 121. Camporesi, E.M., Nielsen, C.H., Bromage, P.R. and Durant, P.A.C., Ventilatory CO2 sensitivity after intravenous and epidural morphine in volunteers, Anesth. Analg. 62, 633–640, 1983.
Ventilatory Effects of Medications Used for Sedation
567
122. Arunasalam, K., Davenport, H.T., Painter, S. and Jones, J.G., Ventilatory response to morphine in young and old subjects, Anaesthesia 38, 529–533, 1983. 123. Daykin, A.P., Bowen, D.J., Saunders, D.A. and Norman, J., Respiratory depression after morphine in the elderly: A comparison with younger subjects, Anaesthesia 41, 910–914, 1986. 124. Bourke, D.L. and Warley, A., The steady-state and rebreathing methods compared during morphine administration in humans, J. Physiol. 419, 509–517, 1989. 125. Peat, S.J., Hanna, M.H., Woodham, M., Knibb, A.A. and Ponte, J., Morphine-6-glucuronide: Effects on ventilation in normal volunteers, Pain 45, 101–104, 1991. 126. Hamunen, K., Ventilatory effects of morphine, pethidine and methadone in children, Br. J. Anaesth. 70, 414–418, 1993. 127. Zhou, H.H., Sheller, J.R., Nu, H., Wood, M. and Wood, A.J.J., Ethnic differences in response to morphine, Clin. Pharmacol. Ther. 54, 507–513, 1993. 128. Lynn, A.M., Nespeca, M.K., Opheim, K.E. and Slattery, J.T., Respiratory effects of intravenous morphine infusions in neonates, infants, and children after cardiac surgery, Anesth. Analg. 77, 695–701, 1993. 129. Houghton, I.T., Aun, C.S.T., Wong, Y.C., Chan, K., Lau, J.T.F. and Oh, T.E., The respiratory depressant effect of morphine: A comparison study in three ethnic groups, Anaesthesia 49, 197–201, 1994. 130. Thompson, P.I., Joel, S.P., John, L., Wedzicha, J.A., Maclean, M. and Slevin, M.L., Respiratory depression following morphine and morphine-6glucuronide in normal subjects, Br. J. Clin. Pharmacol. 40, 145–152, 1995. 131. Borgbjerg, F.M., Nielsen, K. and Franks, J., Experimental pain stimulates respiration and attenuates morphine-induced respiratory depression: A controlled study in human volunteers, Pain 64, 123–128, 1996. 132. Dahan, A., Sarton, E., Teppema, L. and Olievier, C., Sex-related differences in the influence of morphine on ventilatory control in humans, Anesthesiology 88, 903–913, 1998. 133. Sarton, E., Teppema, L. and Dahan, A., Sex differences in morphine-induced ventilatory depression reside within the peripheral chemoreflex loop, Anesthesiology 90, 1329–1338, 1999. 134. Bailey, P.L., Lu, J.K., Pace, N.L., Orr, J.A., White, J.L., Hamber, E.A., Slawson, M.H., Crouch, D.J. and Rollins, D.E., Effects of intrathecal morphine on the ventilatory response to hypoxia, N. Engl. J. Med. 343, 1228– 1234, 2000. 135. Cepeda, M.S., Farrar, J.T., Roa, J.H., Boston, R., Meng, Q.C., Ruiz, F., Carr, D.B. and Strom, B.L., Ethnicity influences morphine pharmacokinetics and pharmacodynamics, Clin. Pharmacol. Ther. 70, 351–361, 2001. 136. Keats, A.S., The effect of drugs on respiration in man, Ann. Rev. Pharmacol. Toxicol. 25, 41–65, 1985. 137. Bellville, J.W., Howland, W.S., Seed, J.C. and Houde, R.W., The effect of sleep on the respiratory response to carbon dioxide, Anesthesiology 20, 628–634, 1959.
568
Gross and Cerza
138. Berkenbosch, A., Bovill, J.G., Dahan, A., DeGoede, J. and Olievier, I.C.W., The ventilatory sensitivities from Read’s rebreathing method and the steadystate method are not equal in man, J. Physiol. 411, 367–377, 1989. 139. Campbell, D., Masson, A.H.B. and Norris, W., The clinical evaluation of narcotic and sedative drugs II: A re-evaluation of pethidine and pethilorfan, Br. J. Anaesth. 37, 199–207, 1965. 140. Downes, J.J., Kemp, R.A. and Lambertsen, C.J., The magnitude and duration of respiratory depression due to fentanyl and meperidine in man, J. Pharmacol. Exp. Ther. 158, 416–420, 1967. 141. Rouge, J.C., Levallorphan and meperidine mixtures: A dose response study of the ventilatory sensitivity to carbon dioxide, Acta Anaesthesiol. Scand. 13, 87–96, 1969. 142. Reier, C.E. and Johnstone, R.E., Respiratory depression: Narcotic versus narcotic-tranquilizer combinations, Anesth. Analg. 49, 119–124, 1970. 143. Kaufman, R.D., Aqleh, K.A. and Bellville, J.W., Relative potencies and durations of action with respect to respiratory depression of intravenous meperidine, fentanyl and alphaprodine in man, J. Pharmacol. Exp. Ther. 208, 73–79, 1979. 144. Rigg, J.R.A., Ilsley, A.H. and Vedig, A.E., Relationship of ventilatory depression to steady-state blood pethidine concentrations, Br. J. Anaesth. 53, 613–619, 1981. 145. Paulson, B.A., Becker, L.D. and Way, W.L., The effects of intravenous lorazepam alone and with meperidine on ventilation in man, Acta Anaesthesiol. Scand. 27, 400–402, 1983. 146. Hersh, E.V., Desjardins, P.J., Simpser, M. and Dadzie, C., Measurement of meperidine-induced respiratory depression using a new non-invasive technique, Anesth. Prog. 32, 194–198, 1985. 147. Flacke, J.W., Flacke, W.E., Bloor, B.C., Van Etten, A.P. and Kripke, B.J., Histamine release by four narcotics: A double-blind study in humans, Anesth. Analg. 66, 723–730, 1987. 148. Kaiko, R.F., Foley, K.M., Grabinski, P.Y., Heidrich, G., Rogers, A.G., Inturrisi, C.E. and Reidenberg, M.M., Central nervous system excitatory effects of meperidine in cancer patients, Ann. Neurol. 13, 180–185, 1983. 149. Harper, M.H., Hickey, R.F., Cromwell, T.H. and Linwood, S., The magnitude and duration of respiratory depression produced by fentanyl and fentanyl plus droperidol in man, J. Pharmacol. Exp. Ther. 199, 464–468, 1976. 150. Stoeckel, H., Schu¨ttler, J., Magnussen, H. and Hengstmann, J.H., Plasma fentanyl concentrations and the occurrence of respiratory depression in volunteers, Br. J. Anaesth. 54, 1087–1095, 1982. 151. Scamman, F.L., Ghonheim, M.M. and Korttila, K., Ventilatory and mental effects of alfentanil and fentanyl, Acta Anaesthesiol. Scand. 28, 63–67, 1984. 152. Negre, I., Gueneron, J., Ecoffey, C., Penon, C., Gross, J.B., Levron, J. and Samii, K., Ventilatory response to carbon dioxide after intramuscular and epidural fentanyl, Anesth. Analg. 66, 707–710, 1987. 153. Bailey, P.L., Streisand, J.B., East, K.A., East, T.D., Isern, S., Hansen, T.W., Posthuma, E.F.M., Rozendaal, F.W., Pace, N.L. and Stanley, T.H., Differences in magnitude and duration of opioid-induced respiratory
Ventilatory Effects of Medications Used for Sedation
154.
155. 156.
157.
158.
159.
160.
161.
162.
163.
164. 165.
166.
167.
569
depression and analgesia with fentanyl and sufentanil, Anesth. Analg. 70, 8–15, 1990. Glass, P.S.A., Iselin-Chaves, I.A., Goodman, D., Delong, E. and Hermann, D.J., Determination of the potency of remifentanil compared with alfentanil using ventilatory depression as the measure of opioid effect, Anesthesiology 90, 1556–1563, 1999. Babenco, H.D., Conard, P.F. and Gross, J.B., The pharmacodynamic effect of a remifentanil bolus on ventilatory control, Anesthesiology 92, 393–398, 2000. Amin, H.M., Sopchak, A.M., Esposito, B.F., Henson, L.G., Batenhorst, R.L., Fox, A.W. and Camporesi, E.M., Naloxone-induced and spontaneous reversal of depressed ventilatory responses to hypoxia during and after continuous infusion of remifentanil or alfentanil, J. Pharmacol. Exp. Ther. 274, 34–39, 1995. Bouillon, T., Bruhn, J., Rand-Radulescu, L., Andresen, C., Cohane, C. and Shafer, S.L., A model of the ventilatory depressant potency of remifentanil in the non steady state, Anesthesiology 99, 779–787, 2003. Fourel, D., Almanza, L., Aubouin, J.P. and Guiavarch, M., Remifentanil: Postoperative respiratory depression after purging of the infusion line, Ann. Fr. Anesth. Reanim. 18, 358–359, 1999. Akiyama, Y., Nishimura, M., Suzuki, A., Yamamoto, M., Kishi, F. and Kawakami, Y., Naloxone increases ventilatory response to hypercapnic hypoxia in healthy adult humans, Am. Rev. Respir. Dis. 142, 301–305, 1990. Santiago, T.V., Sheft, S.A., Khan, A.U. and Edelman, N.H., Effect of naloxone on the respiratory responses to hypoxia in chronic obstructive pulmonary disease, Am. Rev. Respir. Dis. 130, 183–186, 1984. Glass, P.S.A., Jhaveri, R.M. and Smith, L.R., Comparison of potency and duration of action of nalmefene and naloxone, Anesth. Analg. 78, 536–541, 1994. Amin, H.M., Sopchak, A.M., Foss, J.F., Esposito, B.F., Roizen, M.F. and Camporesi, E.M., Efficacy of methylnatrexone versus naloxone for reversal of morphine-induced depression of hypoxic ventilatory response, Anesth. Analg. 78, 701–705, 1994. Snir-Mor, I., Weinstock, M., Davidson, J.T. and Bahar, M., Physostigmine antagonizes morphine-induced respiratory depression in human subjects, Anesthesiology 59, 6–9, 1983. Ramamurthy, S., Steen, S.N. and Winnie, A.P., Doxapram antagonism of meperidine-induced respiratory depression, Anesth. Analg. 54, 352–356, 1975. Randall, N.P.C., Pleuvry, B.J., Fazackerley, E.J., Modla, C.Y., Prescott, L.F. and Healy, T.E.J., Effect of oral doxapram on morphine-induced changes in the ventilatory response to carbon dioxide, Br. J. Anaesth. 62, 159–163, 1989. Cohen, R., Finn, H. and Steen, S.N., Effect of diazepam and meperidine, alone and in combination, on respiratory response to carbon dioxide, Anesth. Analg. 48, 353–355, 1969. Mulroy, M.F., Coombs, J.H.B., Isenberg, M.D. and Fairley, H.B., Age, chronic obstructive pulmonary disease, and InnovarÕ induced ventilatory depression during regional anesthesia, Anesth. Analg. 56, 826–830, 1977.
570
Gross and Cerza
168. Becker, L.D., Paulson, B.A., Miller, R.D., Severinghaus, J.W. and Eger, E.I., II, Biphasic respiratory depression after fentanyl–droperidol or fentanyl alone used to supplement nitrous oxide anesthesia, Anesthesiology 44, 291–296, 1976. 169. Gabuthuler, M.L. and Kaufman, R.D., Respiratory depression of intravenous hydroxyzine in man: Potency, duration, and lack of reversal by naloxone, Anesth. Analg. 60, 634–637, 1981. 170. Zsigmond, E.K., Flynn, K. and Shively, J.G., Effect of hydroxyzine and meperidine on arterial blood gases in healthy human volunteers, J. Clin. Pharmacol. 29, 79–84, 1989. 171. Olson, L.G., Hensley, M.J. and Saunders, N.A., The effects of combined morphine and prochlorperazine on ventilatory control in humans, Am. Rev. Respir. Dis. 133, 558–561, 1986. 172. Zsigmond, E.K., Matsuki, A., Kothary, S.P. and Jallad, M., Arterial hypoxemia caused by intravenous ketamine, Anesth. Analg. 55, 311–314, 1976. 173. Parker, R.I., Mahan, R.A., Giugliano, D. and Parker, M.M., Efficacy and safety of intravenous midazolam and ketamine as sedation for therapeutic and diagnostic procedures in children, Pediatrics 99, 427–431, 1997. 174. Marx, C.M., Stein, J., Tyler, M.K., Nieder, M.L., Shurin, S.B. and Blumer, J.L., Ketamine–midazolam versus meperidine–midazolam for painful procedures in pediatric oncology patients, J. Clin. Oncol. 15, 94–102, 1997. 175. Bourke, D.L., Malit, L.A. and Smith, T.C., Respiratory interactions of ketamine and morphine, Anesthesiology 66, 153–156, 1987. 176. Mildh, L., Taittonen, M., Leino, K. and Kirvela¨, O., The effect of low-dose ketamine on fentanyl-induced respiratory depression, Anaesthesia 53, 965–970, 1998. 177. Mortero, R.F., Clark, L.D., Tolan, M.M., Metz, R.J., Tsueda, K. and Sheppard, R.A., The effects of small-dose ketamine on propofol sedation: Respiration, postoperative mood, perception, cognition, and pain, Anesth. Analg. 92, 1465–1469, 2001. 178. Badrinath, S., Avramov, M.N., Shadrick, M., Witt, T.R. and Ivankovich, A.D., The use of a ketamine–propofol combination during monitored anesthesia care, Anesth. Analg. 90, 858–862, 2000. 179. Gross, J.B. and Long, W.B., Nasal oxygen alleviates hypoxemia in colonoscopy patients sedated with midazolam and meperidine, Gastrointest. Endosc. 36, 26–29, 1990. 180. Vargo, J.J., Zuccaro, G., Dumot, J.A., Conwell, D.L., Morrow, J.B. and Shay, S.S., Automated graphic assessment of respiratory activity is superior to pulse oximetry and visual assessment for the detection of early respiratory depression during therapeutic upper endoscopy, Gastrointest. Endosc. 55, 826– 831, 2002.
15 Central Effects of General Anesthesia
ECKEHARD A.E. STUTH Department of Anesthesiology, Medical College of Wisconsin and Children’s Hospital of Wisconsin Milwaukee, Wisconsin
I.
EDWARD J. ZUPERKU and ASTRID G. STUCKE Department of Anesthesiology, Medical College of Wisconsin Milwaukee, Wisconsin
Introduction
A vast descriptive literature has been accumulated on the respiratory effects of general anesthetics. However, progress in the identification of mechanisms by which anesthetics depress the brainstem respiratory center and alter respiratory rhythm and pattern generation has been slow. This chapter is divided into five parts. Section II summarizes the most important descriptive aspects of the effect of general anesthesia on respiratory rate and depth as reflected in tidal volume, phrenic nerve activity, and respiratory neuron activity. Section III presents the current knowledge of anesthetic effects on fast synaptic neurotransmission as is relevant for respiratory neurotransmission. In section IV an overview of the brainstem respiratory network and the neurotransmission within will be provided. We introduce a prototype respiratory oscillator model that can predict some of the observed effects of anesthetics on respiratory rate. We conclude in section V with three important paradigms that provide an approach to bridge in vitro and in vivo data in order to assess the clinical relevance of the postulated anesthetic mechanisms. This chapter does not attempt to be comprehensive, 571
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Table 15.1
Table of Frequently used Abbreviations
5-HT AMPA AP5 BPM CNS CO2 CSF E EPSP FRC GABA I i.v. IPSP MAC mm Hg nACh receptor NBQX NMDA NTS O2 PACO2 PaCO2 PCO2 PEEP PETCO2 PSR TE TI TTX VT
Serotonin a-Amino-3-hydroxy-5-methylisoxazole-4-propionate 2-Amino-5-phosphonovalerat Breaths per minute Central nervous system Carbon dioxide Cerebrospinal fluid Expiratory Excitatory postsynaptic potential Functional residual capacity g-Aminobutyric acid Inspiratory Intravenous Inhibitory postsynaptic potential Minimal alveolar concentration Millimeter mercury Nicotinic acetylcholine receptor 2,3-Dihydroxy-6-nitro-7-sulfamoylbenzo-(f)quinoxaline N-methyl-D-aspartate Nucleus tractus solitarius Oxygen Partial alveolar pressure for carbon dioxide Partial arterial pressure for carbon dioxide Partial pressure for carbon dioxide Positive end-expiratory pressure Partial endtidal pressure for carbon dioxide Pulmonary stretch receptor Expiratory phase duration Inspiratory phase duration Tetrodotoxin Tidal volume
but it provides a concise update on anesthetic and central respiratory mechanisms and highlights promising areas of respiratory and anesthetic research. We hope it will serve to stimulate further research. Please refer to Table 15.1 for frequently used abbreviations. II.
General Effects of Anesthetics on Respiration
All volatile and most intravenous general anesthetics currently in clinical use cause significant respiratory depression at concentrations suitable for surgery [1–3]. The alveolar concentration of volatile anesthetics can
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be readily measured in real time, so that the applied concentration can be precisely titrated. It is therefore easy to accurately describe the dose-dependent respiratory effects of all inhalational agents in vivo. Traditionally, the potency of inhalational anesthetics has been measured as the minimum alveolar concentration (MAC) [4,5]. The endpoint for MAC determination is lack of purposeful movement to a defined afferent stimulus such as a surgical incision in humans or application of a tail clamp in other mammals. One MAC is defined as the anesthetic concentration where 50% of subjects will not respond with purposeful movement to such stimuli. It has recently been shown that the lack of movement depends on the anesthetic effect on the spinal cord [6], while the anesthetic concentrations in the cortex necessary to suppress such motor responses are much higher [7]. Since the range of MAC for an anesthetic is quite narrow within a species and also across most vertebrate species, MAC remains a useful index to compare anesthetic potency between volatile agents. However, many anesthetic effects on respiration and cortical function occur already in the sub-MAC range (Figure 15.1). For example, it has been well established that the peripheral chemoreflexes in humans are exquisitely sensitive to volatile anesthetics in a range where memory function remains relatively intact (0.1–0.2 MAC) [8,9]. These MACsurgical
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Figure 15.1 Dose-dependent effects of volatile anesthesia on cardiorespiratory and clinical parameters in humans. By convention, 1 MAC refers to 1 MACsurgical, which is the anesthetic concentration at which 50% of humans will not show purposeful movement to skin incision.
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effects are discussed in detail by Dahan et al. (this volume, chapter 16). Similarly, upper airway patency is compromised at sub-MAC concentrations [10]. This has been further quantified in research animals where both the efferent nerves and the muscles responsible for upper airway patency were significantly depressed by volatile anesthetics at about 0.5 MAC [11–13] while the pump muscles, in particular the diaphragm, still generated adequate force. The selective depression of upper airwayrelated respiratory motoneuron groups may be due to the presence of anesthesia-activated potassium leak channels [14], which will be discussed below in sections III.A and V.C. Remarkably, the rhythm-and pattern generating neurons in the brainstem respiratory center as well as the major respiratory pump muscle, the diaphragm, show a relative resilience to anesthetic depression. A. Anesthetic Effects on Respiratory Minute Ventilation Effects of Volatile Anesthetics
Humans and other mammals typically maintain adequate minute ventilation at surgical concentrations of volatile anesthesia provided the airway is kept patent (Figure 15.2). This is only possible because of a major increase in the arterial CO2 partial pressure. Anesthetics greatly depress the respiratory drive, which is expressed as a decrease in tidal volume. The concomitant arterial CO2 (PaCO2 ) increase compensates in part for the anesthetic depression of drive, preserving spontaneous alveolar minute ventilation at 1 MAC of most volatile anesthetics at a magnitude comparable to the awake state. In the absence of anesthesia, minute ventilation approximately doubles when PaCO2 is elevated by as little as 2 mm Hg [15]. The relationship is linear in the moderate hypercapnic range (40–70 mm Hg) [1]. The increase of PaCO2 caused by 1 MAC volatile anesthesia is in the range of 5–15 mm Hg depending on the anesthetic [1,16]. The relative respiratory depression for each volatile anesthetic can be gauged from the PaCO2 at which spontaneous respiration occurs. At 1 MAC the depression is greatest, i.e., the PaCO2 is highest for enflurane [17,18] 4desflurane [19] isoflurane [20,21] 4sevoflurane [22,23] halothane [16] (see Figure 15.2). Dahan et al. have compared the effects of subanesthetic concentrations of different anesthetics on respiration with the help of dynamic end-tidal forcing of alveolar gases. This methodology allows differentiation between effects on the central and peripheral chemoreflex loops [24,25] (this volume, chapter 16). In humans the CO2 drive at 1 MAC of modern volatile anesthetics is entirely due to the central CO2 chemoreflex loop, since it has been convincingly shown that the peripheral chemoreflex loop is completely suppressed at 1 MAC of volatile anesthetics [26,27]. Therefore, any changes in the CO2 response at these anesthetic concentrations reflect effects
Central Effects of General Anesthesia Respiratory Rate
40
575 800
H
700
Tidal Volume E
H: halothane E: enflurane
30 H D
ED
H
(ml)
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400 D
S
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I: isoflurane S: sevoflurane D: desflurane
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E IS HI
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10 E
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8 (l/min)
6
H E
H S
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I
40 D
2 Awake
1 MAC
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I S D
4
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70
H I
H
S
H IS H H
D S IH EH
Awake
1 MAC
2
Figure 15.2 Effects of volatile anesthesia on clinical respiratory parameters in humans. (The data are compiled from studies by Refs. 18, 19, 21, 22, and 336 and modified after Ref. 3.)
on the mechanisms of central chemoreception, the respiratory rhythm and pattern generator in the brainstem, and the transmission of inspiratory and expiratory drive to the airway and spinal cord respiratory neurons. The CO2 response curves show that volatile anesthetics raise the apneic threshold and also depress the slope of the CO2 response dose-dependently [1,3]. Such descriptive data allow comparison of the various agents but they do not offer further insight into the mechanisms of central respiratory depression.
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The clinical observations have been confirmed in animal experiments where reflex loops can be opened and the mechanisms of respiratory depression can be further investigated. For example we found in paralyzed, vagotomized dogs that halothane, isoflurane and sevoflurane dosedependently depressed the peak amplitude of the phrenic neurogram (neural tidal volume) [28–30], which is an index of the respiratory tidal volume [31], under hyperoxic isocapnic conditions. In addition, these agents dose-dependently depressed the slope of the CO2 response of the peak phrenic activity [28,29]. The synaptic mechanisms of the depression of respiratory premotor neuron activity in this canine model are described in section V.A. Effects of Intravenous Anesthetics
Intravenous anesthetics have also been shown to decrease respiratory minute ventilation, again mainly by decreasing tidal volume and depressing responsiveness to CO2 with increasing depth of anesthesia. However, unequivocal quantitative data between anesthetic dose and effect on respiration are generally lacking for intravenous agents, because steady-state anesthetic plasma concentrations cannot yet be measured in real time. Propofol
The best human data are available for propofol infusions, where titrated adjustments of infusion rates either manually or by computer can achieve fairly constant plasma concentrations, albeit with large inter-individual variability. The data for propofol indicate that in humans, rib cage ventilation is better preserved with light propofol anesthesia [32] than with inhalational anesthesia [33]. Propofol anesthesia at typical infusion rates to maintain a state of light general anesthesia (6–12 mg kg1 h1) in humans depressed minute ventilation by about 40%, which was due to an approximately 60% decrease in tidal volume and an increase in respiratory rate [34]. Propofol also shifts the CO2 response curve to the right [34]. Even sedative doses of propofol depress the central but not the peripheral chemoresponse to CO2 by about 40% [35]. In spontaneously breathing patients without airway support, sedative doses of propofol cause a phase shift in respiratory pattern between abdomen and rib cage suggesting upper airway obstruction [36,37]. Ketamine
Ketamine is an intravenous anesthetic that provides dissociative anesthesia [38]. An induction bolus of 1 mg/kg, followed by infusion rates of 100–200 mg kg1 min1, provides adequate analgesia for major surgery with maintained spontaneous ventilation [39]. An induction bolus of
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2 mg kg1, followed by an infusion rate of 41 21 mg kg1 min1 and supplemented with 65% nitrous oxide, provides adequate anesthesia with a mean plasma concentration of 9.3 0.8 mmol 1 [40]. Hamza et al. showed in children that a similar regimen (2 mg/kg bolus followed by 40 mg kg1 min1) caused unconsciousness but did not affect baseline respiratory rate, tidal volume (VT), end-tidal CO2 tension, or minute ventilation [41]. However, the ventilatory response to CO2 as expressed by the slope of minute ventilation/end-tidal CO2 relationship was significantly depressed 5 min after the bolus. After 30 min of continuous infusion this slope had returned to baseline, but the CO2 response curve remained shifted to the right indicating a moderate respiratory depression [41]. A similar depression of the CO2 response without change in CO2 sensitivity was observed by Bourke et al. [42] who gave five logarithmically scaled doses of ketamine totalling 3 mg kg1 to volunteers and showed a right shift of the CO2 response of 2.0 1.2 mm Hg. Thus ketamine is the only general anesthetic that does not depress the slope of the CO2 response curve after loss of consciousness. Similar respiratory changes, i.e., a rightward shift without slope change in the CO2 response curve, are also characteristic for opioids in the analgesic dose range as long as the subjects stay awake. This suggests that the respiratory depressant effects of ketamine may in part be mediated via opioid receptors. This has been recently confirmed in m-opioid receptor knockout mice [43]. Large doses of ketamine, which are required for anesthesia in rodents, reduced the hypercapnic response in wild-type mice significantly more than in m-receptor knockout mice, and the opioid antagonist naloxone only reversed the respiratory depressant effect of ketamine in wild-type but not in knockout mice [43]. Interestingly, the breathing pattern of children receiving a ketamine infusion was distinctly different from children receiving comparable halothane anesthesia (1–1.3 MAC) [44]. Ketamine caused a prolongation of the inspiratory duration (TI), i.e., an apneustic type of breathing pattern (Figure 15.3). Expiratory braking was frequently observed. In contrast, halothane decreased TI and increased the respiratory rate. However, the ratio of VT/TI (a measure of mean inspiratory flow rate) was similar between the two agents, as was the respiratory minute ventilation [44]. Similar but more pronounced effects of ketamine are observed in cats [45], which require larger doses for surgical anesthesia than humans or dogs. In cats, 40 mg/kg intraperitoneal ketamine decreases minute ventilation by about a third, while the resting end-tidal CO2 rises by about 5 mm Hg. The breathing pattern is apneustic with a 300% increase in tidal volume, a decrease in respiratory rate, and prolongation of TI from 0.44 to 0.87 of the respiratory cycle. The greatest flow occurs early in inspiration. The apneustic breathing pattern may be responsible for the preservation of the CO2 sensitivity, since increasing CO2 concentration shortens TI without
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A Volume (ml) Respiratory cycle TE
TI Flow (ml/sec)
Time to maximal inspiratory flow
B
Volume (ml)
TI
TE
Flow (ml/sec) 1 sec Time to maximal inspiratory flow
Figure 15.3 Breathing pattern of two children at comparable depths of anesthesia with ketamine (A) and halothane (B). Ketamine anesthesia results in a longer respiratory cycle with prolonged inspiratory phase duration (TI) and larger tidal volume (VT). Note that inspiratory flow rapidly reaches its maximum with ketamine and inspiratory duration is long. TE: expiratory phase (adapted from Ref. 44).
reducing VT, thereby increasing minute ventilation [45]. The apneustic breathing pattern seems to result from an interaction of ketamine with NMDA receptors involved in the central inspiratory off-switch mechanism in a variety of mammals [46–48]. The effect cannot be antagonized by the opioid antagonist naloxone [49], but apneustic respiration can be induced by systemic administration of a variety of NMDA-antagonists such as MK-801 [50]. In humans, ketamine also leads to delayed expiratory emptying, which contributes to maintaining the functional residual capacity (FRC) at levels similar to the awake state [51–53]. Indeed ketamine is the only general anesthetic in clinical use that does not lead to a decrease of FRC. This appears to be related to a sparing of intercostal muscle activity
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during inspiration and a relative increase in rib cage contribution to tidal breathing [54], in contrast to inhalational anesthetics [33,55,56]. It has also been suggested that during ketamine anesthesia, some inspiratory muscle tone is maintained during expiration and more laryngeal braking is preserved when compared with inhalational agents. This may also help to maintain FRC. In conclusion, it appears that anesthetic doses of ketamine affect the control of respiration in two ways: they block central glutamatergic mechanisms mainly involved in inspiratory timing and they depress the central chemodrive response via opioid receptor agonism. The peripheral hypoxic chemoresponse is only modestly depressed in dogs as compared with other intravenous (barbiturates) [57] and inhalational agents [58]. Barbiturates
Gautier et al. described the respiratory effects of barbiturates in otherwise healthy humans who were comatose secondary to barbiturate intoxication [59]. These patients showed an increased respiratory rate (mean 25 breaths/min), shortened respiratory duty cycle, small tidal volume, and reduced minute ventilation. As they slowly awakened from coma over several days, respiratory rate decreased, respiratory duty cycle increased and minute ventilation increased. Tidal volume more than doubled, and the mean inspiratory flow (VT/TI) doubled [59]. Launois et al. reported similar findings in a later series [60]. However, quite different effects are reported for spontaneously breathing cats under deep barbiturate anesthesia; they showed an apneustic breathing pattern with a marked decrease in respiratory rate but relative preservation of tidal volume. The decrease in resting ventilation led to a significant increase in resting alveolar PCO2 [61]. The slope of the ventilatory response to CO2 and the response to hypoxia were markedly depressed by surgical pentobarbital anesthesia in cats compared to the awake state [62]. Similarly, in humans a small induction dose of thiopental significantly decreased the slope of the CO2 response curve [63]. Other species such as neonatal rats show a decrease of respiratory rate with barbiturate anesthesia due to an increase in expiratory time with no change in inspiratory time [64]. The induction of apneusis in some mammals but not humans suggests that barbiturates affect the inspiratory off-switch mechanism in the respiratory rhythm-generating center (see also section IV.B). In cats, neurons responsible for this off-switch receive NMDA receptor-mediated excitation and GABAergic inhibition [65]. Barbiturates enhance GABAA receptor function [66–68]. In addition, inhibition of NMDA receptor function by barbiturates was shown in neurons [69] and transfected oocytes [70]. However, the divergent effects of barbiturates on respiratory rate and pattern in animals and humans suggest that respiratory
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network models based on animal experiments may not be easily extrapolated to man [71]. B. Anesthetic Effects on the Respiratory Rate
All volatile anesthetics depress respiratory chemodrive inputs and thereby tidal volume and minute ventilation, but paradoxically respiratory rate is higher at 1 MAC compared with the awake resting state [3,21,72–76]. In humans, respiratory rate increases beyond 1 MAC with most volatile agents (Figure 15.2), while the effects in animals are more variable [29,77,78]. It has been suggested that this rate increase is due to effects on slowly adapting pulmonary stretch receptors (PSRs, also see section IV.C.2) and pulmonary receptors with unmyelinated fibers (vagal C-fibers) [79] because deafferentation of the lungs by vagotomy significantly blunts the tachypneic effect. Still, in most mammals an increase in respiratory rate can be observed even after vagotomy [29,73,80]. In a feline artificial brainstem perfusion model where halothane can be delivered selectively to the brainstem or the periphery, the anesthetic-induced tachypnea was found to be exclusively of central (brainstem) origin [74]. Using a decerebrate cat model, Gautier et al. suggested that halothane-induced tachypnea was of suprapontine origin, as decerebration appeared to abolish the response [80]. In a vagotomized decerebrate canine model, however, our group still consistently observes an increase in respiratory rate with halothane and sevoflurane [30,81]. In section IV.B.1., we present a functional model of a reciprocal inhibitory oscillator with membrane characteristics imitating those of brainstem respiratory neurons. Simulations show that the hypothetical neuronal network will oscillate faster when excitatory drive is reduced. In agreement with these model predictions, we frequently observe an increase in neural respiratory rate when the CO2 drive is reduced from normocapnia to hypocapnia in our vagotomized canine model. Occasionally we observe a hypocapnic polypnea (i.e., very rapid shallow breathing) just before the apneic threshold is reached. With a further reduction of drive, the oscillator can be locked in the expiratory phase (apnea), e.g., in dogs and cats [28,82], or in the inspiratory phase, e.g., in rabbits [82]. In contrast, an increase in excitatory chemodrive into the hypercapnic range does not result in a significant increase in respiratory rate beyond that seen during normocapnia. The oscillator model also provides a possible explanation for the decrease in respiratory rate with intravenous anesthetics such as barbiturates that has been observed in some mammals (e.g., cats and rabbits) but not humans [71,83,84] (see IV.C.2). An increase in inhibition, which is to be expected if the main anesthetic effect is an enhancement of GABAA receptor function, leads to a slowing of the oscillator. However, many general anesthetics seem to both depress excitatory and enhance inhibitory
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neurotransmission (see section III.B) and the compounded effects on the respiratory oscillator will determine in which direction respiratory rate changes. C. Anesthetic Effects on Respiratory Neurons
The activity of single respiratory neurons in vivo is dose-dependently depressed by all volatile anesthetics (see Figure 15.4A and Refs. [28–30,
Neuron activity, normalized to 0 mac (d-j) or 1 MAC (a-c) (%)
A 120
100
IHH
HS
I H H
(g+h) HS(e+f) (d+f) HS (i+j)
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H I(a)
H: halothane I: isoflurane S: sevoflurane Inspiratory neurons Expiratory neurons
40
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H
H(b) H(b)
1.5
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Figure 15.4 Effects of the volatile anesthetics halothane (H), isoflurane (I), enflurane (E), and sevoflurane (S) on the neuronal discharge activity of respiratory neurons in animals without the use of background anesthesia. (A) Shows data from inspiratory (solid line) and expiratory (dashed line) premotor neurons in the caudal VRG in neuraxis-intact (a–c) and decerebrate dogs (d–j). (a) refers to Ref. 28; (b) to Ref. 29; (c) to Ref. 93; (d) to Ref. 81; (e) to Ref. 30; (f) to Ref. 94; (g) to Ref. 95; (h) to Ref. 86; (i) to Ref. 96; and (j) to Ref. 88. The data are normalized to the neuronal discharge frequency at 0 MAC (decerebrate animals) or 1 MAC anesthesia (intact animals). (B) Shows data from inspiratory neurons in the region of the nucleus ambiguus of the VRG in cats. In two studies the authors compared the effects of halothane and sevoflurane (k ¼ Ref. 85) and halothane and enflurane (l ¼ Ref. 89) with crossover administration of the two agents while recording from the same neuron. The neuronal discharge activity was normalized to the frequency at 1 MAC halothane for each study. (C) Compares the effects of halothane (m ¼ Ref. 90) and enflurane (n ¼ Ref. 91) on inspiratory neurons in the DRG in decerebrate cats. The data are normalized to the neuronal discharge activity at 0 MAC for both studies.
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Neuron activity, normalized to 1 MAC halothane
B 300 S 250 H 200 E H 150 S H
100
H 50
H
E
H H
E
0 0.0
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H E(I) S(k) 1.5
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100 H: halothane E: enflurane
80
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40
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20 H(m) 0 0.0
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1.0
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MAC
Figure 15.4
Continued
76,81,85–95]) and the few intravenous anesthetics that have been tested. Only a few studies have attempted to resolve the mechanisms by which this anesthetic depression of neuronal activity occurs. The studies by Tabatabai were performed on inspiratory neurons in the dorsal respiratory group (DRG) of decerebrate cats (Figure 15.4C) [90,91], and it is likely that most of these neurons are inspiratory premotor neurons providing drive to phrenic motoneurons. The studies by Stuth et al. were performed on
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inspiratory and expiratory premotor neurons in the ventral respiratory group (VRG) of both intact [29,77,93] and decerebrate dogs [30,81,86,88,94–96]. The studies by Kasaba et al. were performed on inspiratory neurons in the nucleus ambiguus of cats, and it is possible that many of these neurons are vagal motoneurons [85,89] (Figure 15.4B). While all agents caused a dose-dependent depression, it seems that the depression occurs at lower MAC levels in decerebrate vs. intact animals, suggesting that decerebration per se may remove some excitatory inputs to the respiratory centers in the brainstem. Three paradigmatic models [93] that will explore the mechanisms by which general anesthetics depress respiratory neurons [30,91,92,94,95,97–100] will be discussed in further detail in section V. III.
Anesthetic Effects on Fast Synaptic Neurotransmission
It has been proposed that there are many targets of anesthetic actions at multiple levels of neuronal networks, and a strong argument can be made that, in particular, volatile anesthetics affect multiple sites of a network simultaneously [101]. Even though at clinically relevant concentrations (0.5–2 MAC) some effects appear so minor that their importance is questionable, it must be considered that the anesthetic effect may result from a cascaded effect on many targets. For many intravenous anesthetics, clinically relevant concentrations are more difficult to establish, since many of these drugs are only used for induction of unconsciousness or in combination with other intravenous drugs, in particular opioids. It is important to realize that the free concentration of intravenous anesthetics in the aqueous phase is relevant for the clinical effect (Table 15.2), and thus receptor action, and not the total plasma concentration, which is often much larger because of high protein binding. For example, propofol is about 98% plasma bound and the MAC equivalent has been estimated as a free aqueous concentration of 1.5 mM [102]. Some authors suggest that only even lower concentrations of propofol (0.4 mM) are clinically relevant [103]. This brief review will focus on molecular targets that are affected at clinically relevant concentrations. There is overwhelming evidence that synaptic neurotransmission, especially fast synaptic neurotransmission via ligand-gated ion channels, is of central importance for information processing in neural networks. This is also true for the brainstem respiratory network (see section IV). A large amount of data shows that the mechanisms involved in synaptic neurotransmission are markedly affected by clinical concentrations of volatile and intravenous anesthetics. For an up-to-date review we refer to an entire issue of the British Journal of Anesthesia (Vol. 89(1), 2002). In the
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Table 15.2 Anesthetic Halothane Isoflurane Enflurane Sevoflurane Desflurane Thiopental Pentobarbital Ketamine Propofol Etomidate
Aqueous Concentrations of General Anesthetics in Humans Partial pressure (humans) (% atm)
EC50 free plasma concentration (mM)
0.75 1.30 1.70 2.05 6.00
190 270 490 300 530 14 88 15 1.7 2.4
Data are given for the EC50 of intravenous agents and 1 MAC of volatile agents, adapted from Refs. 337 and 156.
following subsections we will present a brief overview of the anesthetic effects on voltage-gated and ligand-gated ion channels, which mediate presynaptic neurotransmitter release and postsynaptic neuronal excitability. A. Anesthetic Effect on Voltage-gated Ion Channels
Voltage-gated channels are considered to be relatively insensitive to clinical concentrations of anesthetics. However, because of the dominant role of calcium [104–109] in presynaptic neurotransmitter release, it has been argued that cascaded effects may render even modest depression of these channels significant for neuronal function. Action potentials propagated by sodium currents depolarize the membranes of synaptic terminals, opening voltage-gated calcium channels. Extracellular calcium ions enter the nerve terminal and trigger the release of neurotransmitters (Figure 15.5). The main transmitters in brainstem respiratory neurons are glutamate and GABA. Studies in synaptosomes (isolated nerve endings that behave like presynaptic terminals) have shown that volatile anesthetics can depress glutamate secretion [110]. There is also evidence that GABA secretion can be inhibited by anesthetics [111,112]. Studies in spinal cord motoneurons that used quantal analysis showed that halothane and barbiturates depressed synaptic neurotransmission by decreasing action potential-evoked secretion of neurotransmitters [113,114]. Further evidence that volatile anesthetics depress glutamatergic transmission via presynaptic actions was derived from hypothalamic slices [115].
Central Effects of General Anesthesia AP4 BLOCKS = DEPOL
VERATRADINE ACTIVATES
Na+
AP
585
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Ca++ K+
1 IONOMYCIN=IONOPHORE
2 1
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Anesthetic block No anesthetic effect
1,2
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Anesthetic reduction Ca++ 1: Schlame and Hemmings, 1995 2: Miao et al., 1995
1,2
Figure 15.5 Steps involved in presynaptic neurotransmitter release and anesthetic effects on these mechanisms. Action potentials (AP) travelling down the axon lead to opening of voltage-gated Naþ channels and to a depolarization of the membrane of the synaptic bouton. This depolarization leads to an opening of voltage-gated Ca2þ channels, which results in an increase in intracellular calcium ([Ca2þi]). Calcium binds to neurotransmitter containing vesicles, which mediates the transmitter release. This process can be experimentally triggered at every step of the cascade (encased), i.e., by AP4-block of KA channels, which leads to depolarization, by direct activation of Naþ channels by veratridine; by depolarization of the membrane with KCl, which leads to opening of voltage-gated Ca2þ channels, or by directly allowing calcium influx with the ionophore ionomycin. Two studies have shown that halothane [110,139], enflurane, and isoflurane [139] depress glutamate release from synaptosomes by interfering with parts of this cascade (see text for details).
It has been suggested that the neurosecretory mechanism does not significantly differ for various types of neurotransmitters and that it is therefore reasonable to assume that anesthetics would affect presynaptic neurotransmitter release similarly for inhibitory and excitatory synapses [116]. However, Westphalen and Hemmings have recently shown in synaptosomes that isoflurane depressed glutamate release more than GABA release [117]. Independent of anesthetic effects on neurotransmitter release, it is clear that any anesthetic-induced depression of the activity of presynaptic neurons per se will lead to decreased action potential firing and thus decreased neurotransmitter release, as demonstrated in hippocampal slices [118].
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Voltage-gated calcium channels are found in all types of respiratory neurons [119] but there are no studies to date on anesthetic effects on these channels in respiratory neurons. At least six types of voltage-gated calcium channels are known in the mammal [105,120–123]. T- [124], L-, N- and P-type calcium channels have been extracted from the brain. T-type channels activate at a more negative membrane potential than the other channels, i.e., they are low-voltage activated (LVA) and rapidly inactivate. They have been implicated in pacemaker activity [125] and repetitive firing of neurons [126], and can be potently depressed by anticonvulsants [127]. L-type channels are high-voltage activated (HVA), inactivate slowly, and are fairly ubiquitous [128], also occurring in respiratory neurons [129]. They appear to be involved in the regulation of neuronal excitability [130] but do not contribute to synaptic neurotransmitter release [131]. P- and N-type channels are HVA and inactivate at a slower rate than the T-type channels. P-type channels may constitute as much as 80% of the voltage-gated calcium channels in mammalian CNS nerve terminals, and they have been linked to stimulus secretion coupling [132]. P-type channels were initially found in cerebellar Purkinje cells and are potently and selectively depressed by the spider toxin omega-AGA-IVA [133]. N-type channels were initially found in peripheral neurons and appear to be specific for neurons and some secretory cells. N-type channels are selectively depressed by the snail toxin omega-conotoxin-GVIA [132] and have also been linked to stimulussecretion coupling of transmitters [134]. Hippocampal pyramidal neurons exhibit T-, L-, N-type and a 4th calcium current (possibly P-type). Isoflurane inhibited both high and low voltage-activated calcium currents in these neurons with an inhibition of 50% at about 1.3 MAC isoflurane for the sustained current and at about 2.6 MAC isoflurane for the peak current [135] (Table 15.3). Since voltagegated calcium channels at synaptic terminals are thought to open only for a very short time (about 1 msec [136]) in response to an action potential, it has been suggested that the peak currents are more relevant for neurotransmitter release [137]. P-type calcium channels were also studied in Purkinje cells and found to be relatively insensitive to a variety of general anesthetics at clinically relevant concentrations. Inhibition by about 1.3 MAC halothane, 1.3 MAC isoflurane, 32 mM thiopental, 50 mM pentobarbital, and 2 mM propofol was 10% [137]. Based on these findings, Franks and Lieb claim that anesthetic effects on calcium channels are clinically irrelevant [102]. However, Lynch and Pancrazio have argued that even small effects on calcium channels may have clinical relevance because the synaptic release of neurotransmitters is a high-order function of the calcium that enters during depolarization of the presynaptic terminal [138]. For example, they calculate that a 15% anesthetic reduction of the control
Anesthetic Halothane
Effects of General Anesthetics on Voltage-gated Ion Channels TTX-sensitive Naþ channel
P-type Ca2þ channel
ø: [338]a I: [140a,141b], [110,142]c I: [141 ], [144] ø: [110]
ø: [110,137] I: [139] ø: [137,339,144] I: [135,139] ø: [110,137]
I: [143,144]
ø: [137,144]
Isoflurane b
Barbiturates Ketamine Propofol
T,L,N-type Ca2þ channel
Voltage-gated Kþ channel
SKCa Kþ channel
ø:[153], [155] I: [110 ] ø: [339] I: [135]
ø: [155]d
ø: [157] ø: [157] ø: [157]
ø: [144,340]
ø: [156] ø: [153,156] ø: [156]
Central Effects of General Anesthesia
Table 15.3
Included are channels involved in neuronal presynaptic transmitter release, postsynaptic neuronal excitability or axonal conduction. Included are studies where receptors have been expressed in non-neuronal cells. Only studies that used anesthetic concentrations in the clinical range, i.e., 52 MAC for volatile anesthetics and 52 ED50 for intravenous anesthetics, are listed. Anesthetic effects are only considered relevant when they cause a 10% change from baseline channel function (compare Ref. 102). (ø: no effect on channel function; I: inhibition of channel function). a Channel located on axons. b IIA subtype of the rat brain sodium channel. c Channel located on presynaptic nerve endings. d Experiment conducted with methoxyflurane.
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calcium current to 85% may be amplified to a 39–48% depression of transmitter release (cascaded effects: (0.85)3 ¼ 0.61 or (0.85)4 ¼ 0.52 [138]). The presynaptic nerve terminal is not accessible for direct examination or recording, however synaptosomes can be used to study presynaptic neurotransmitter release. Depolarization of synaptosomes through block of Kþ channels results in an activation of TTX-sensitive Naþ channels, a depolarization of the membrane, which leads to an increase in intrasynaptosomal calcium and subsequent exocytosis of glutamate (see Figure 15.5). This sequence requires extracellular calcium, suggesting that transmitter release involves calcium entry rather than calcium release from intracellular stores [110,139]. Studies in cerebral synaptosomes showed that 1–2 MAC isoflurane, halothane or enflurane dose-dependently decreased the KCl-induced increase in both [Ca]i and glutamate release by about 10–20% per MAC [139]. The decrease in [Ca]i was sufficient to account for the associated decrease in glutamate release. However, in a similar preparation, Schlame and Hemmings found that volatile anesthetics also reduced the increase in [Ca]i and glutamate release when the synaptosomes were depolarized by activation of the TTX-sensitive Naþ channels [110]. In their preparation, anesthetics depressed the increase in [Ca]i when the synaptosomes were depolarized with KCl, i.e., when voltage-gated calcium channels were directly activated, but glutamate release was not reduced [110]. Both groups concluded that volatile anesthetics did not interfere with the mechanism by which calcium caused exocytosis of the neurotransmitter but suggested that the anesthetics may have inhibited voltage-dependent calcium channels. Schlame and Hemmings, however, argued that the lack of decrease in glutamate release after KCl depolarization indicated that the calcium channels blocked by the anesthetic were not related to exocytosis. They suggested that anesthetic inhibition of voltage-gated sodium rather than calcium channels may be the relevant mechanism for the depression of glutamate release [110]. Voltage-gated Sodium Channels
Axonal conduction, which depends on action potential propagation via sodium channels, is quite insensitive even to pharmacologic doses of general anesthetics [140]. However, it has recently been shown that the IIA subtype of the rat brain sodium channel, which was expressed in Chinese hamster ovarian cells, showed a strong voltage-dependent anesthetic depression with a 50% depression of current at 1 MAC when the resting potential was held at 60 mV [141]. The relevance of these anesthetic effects on Naþ channels for general anesthesia remains unclear, because it is known that more than 70% of all Naþ channels have to be blocked to prevent action potential firing in axonal conduction [141].
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Hemmings’ group has recently shown in synaptosomes that Naþchannels involved in neurotransmitter release were also moderately sensitive to halothane at 1.5 MAC [142] and to propofol [143]. In a subsequent study, they found that 2 MAC isoflurane depressed glutamate release evoked through Naþ-channel activation but not evoked through Ca2þchannel activation [144]. However, the effects on the same mechanisms were negligible for propofol at clinically relevant concentrations. The authors argue that even relatively small changes in sodium channel conduction can produce large shifts in the equilibrium potential with profound functional consequences. Potassium Channels
Kþ channels have been organized into four families: the voltage-gated Kþ channels, Kv; the calcium-activated Kþ channels [145], KCa; the inward rectifier Kþ channels [146], KIR, and the tandem pore domain Kþ channels. For a detailed review, we refer to Yost’s article [147]. The much larger concentration of Kþ ions on the inside of the cell membrane vs. outside is the main determinant of the resting membrane potential of excitable cells such as neurons. The resting potential is in the range of 50 to 70 mV for most neurons. Neuronal Kþ channels are responsible for repolarization of the neuronal cell membrane after an action potential [148]; they cause afterhyperpolarization [149] of neurons, and they are important for postsynaptic temporal integration of neuronal signals [150]. Kþ-channels limit action potential duration, determine spike frequency in neurons [151] and serve to protect the neuron from excessive excitation [149]. Since Kþ-channels are fundamentally important in governing the excitability of neurons, modulation of Kþ-channel activity by anesthetics could be involved in the global CNS depression produced by these agents. Any drug that increases the opening of a particular Kþ channel will hyperpolarize the cell and decrease its excitability. In contrast, agents that depress the outward Kþ current can lead to an increase in neuronal excitability. For example, in the rare familial syndrome of episodic ataxia, a point mutation of the Kv1.1 gene leads to a defective voltage-gated Kþ channel in neurons. Individuals with this mutation suffer from episodic neuronal hyperexcitability, which leads to ataxia [152]. Anesthetic insensitivity of voltage-gated potassium channels
So far, voltage-gated Kþ channels, which contribute to the action potentials of the neurons, have been found to be relatively insensitive to all types of general anesthetics [153–155] (Table 15.3). The effects of thiopental, pentobarbital, methohexital, propofol, ketamine, midazolam, and droperidol on neuronal Shaw-like voltage-dependent Kþ currents of human origin were recently studied in SH-SY5Y human neuroblastoma cells [156]. These
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Kþ currents have a high threshold of activation and a high sensitivity to tetraethylammonium and 4-aminopyridine similar to Shaw (Kv3) Kþ channels and are found in the auditory system and the hippocampus. The IC50 values (half-maximal inhibition) ranged from 100 to 1000 times the clinically relevant plasma concentration [156]. Kþ-current suppression at clinically relevant aqueous free plasma concentrations (Table 15.2) was estimated to be only 3% for the anesthetic agents. Nonetheless the authors suggest that this small in vitro effect may be clinically relevant [156], although others clearly disagree with this view [102]. Small-conductance Kþ channels (SK) are part of the afterhyperpolarization and prevent the neuron from hyperexcitability [149]. A study on recombinant SK2 channels expressed in human embryonic kidney (HEK 293) cells showed that clinical concentrations of volatile anesthetics, ketamine and methohexital did not alter the channel conductance [157]. The conductance of inward rectifier potassium channels (KIR) is strongly voltage-dependent and regulates the membrane potential and thus overall excitability of the cell [146]. Some of the KIR channels are coupled with a G-protein, i.e., binding of a neurotransmitter activates the channel leading to hyperpolarization of the membrane [158]. Interestingly, opioids exert their analgesic effects by binding to opioid receptors, which are G-protein coupled, and thus lead to opening of Kþ channels and neuronal hyperpolarization [159]. Tandem pore potassium channels
The fourth Kþ-channel family is tandem pore domain Kþ channels [160]. These are not voltage-gated and are responsible for a baseline or leak conductance. These channels are unique because they contain two Kþ-selective central pores in tandem. Tandem pore channels are believed to be by far the most abundant Kþ channels. Members of this family have also been implicated in oxygen sensing in the carotid body [161]. Another member of this family, the TASK-1 (Tandem pore Acid Sensitive Kþ) channel, was investigated in hypoglossal motoneurons (HMs) where it was inhibited by multiple neurotransmitters [162]. Some members of the tandem pore Kþ channels are activated by clinically relevant concentrations of volatile anesthetics [14,163,164], which causes hyperpolarization of the neurons and thus decreases their excitability (see also section V.C). Several earlier studies showed that various volatile anesthetics (ether, halothane, and isoflurane) could hyperpolarize certain CNS neurons as a result of an increased Kþ conductance [165,166]. Similarly, central pacemaker neurons from the pond snail ceased firing when hyperpolarized by volatile anesthetics at clinical concentrations [167]. Only anesthetic-sensitive snail neurons possessed a unique anesthetic-activated outwardly rectifying Kþ current, which was not voltage-gated and did not desensitize. The current appeared to be related
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to background potassium currents, which was subsequently confirmed [168]. A similar type of Kþ current is activated by halothane in rat motoneurons [100]. Recently, Patel et al. reported that recombinant mammalian (mouse and human) TREK-1 (TWIK-1-Related Kþ-channel 1) and TASK channels, which belong to the tandem pore family, e.g., TREK-1, TRAAK, TWIK-1 (Tandem Pore Weak Inward rectifying Kþchannel-1) and TASK-1, were dose-dependently activated by halothane and isoflurane and that significant Kþ-channel opening already occurred at MAC concentrations [14]. TREK-1 and TASK channels are particularly abundant in the brain. It has been argued that a large number of Kþ-background channels are yet to be discovered in man since they are the most abundant Kþ channels found in invertebrates (C. elegans) and that they will likely be found to be highly relevant for explaining general anesthetic effects [163]. B. Anesthetic Effect on Ligand-gated Ion Channels
Fast synaptic neurotransmission depends on the secretion of neurotransmitters that bind to ligand-gated ionotropic receptors, which then activate ion channels. The most important excitatory neurotransmitter in the respiratory network is glutamate [169,170] that activates the AMPA/kainate (a-amino-3-hydroxy-5-methylisoxazole-4-propionic acid) receptor and the NMDA (N-methyl-D-aspartate) receptor. Another important excitatory neurotransmitter is acetylcholine acting on nicotinic acetylcholine (nACh) receptors [171]. Postsynaptic ligand-gated ion channels have been proposed as the principal targets of anesthetic action [172,173]. Glutamate Receptors
In early studies in cerebral slices, halothane seemed to have little depressant effect on glutamate receptors [174,175]. Other studies showed that NMDA receptor-mediated responses in cortical [176], hippocampal, and spinal cord neurons were depressed by dissociative anesthetics such as ketamine and phencyclidine [177–179]. NMDA receptor depression was also reported with enflurane [180], halothane [181], isoflurane [182,183] and barbiturates [70], but propofol was ineffective at clinically relevant concentrations [184] (Table 15.4). The AMPA receptor-mediated component was depressed by isoflurane but not by ketamine [179], barbiturates or propofol [185]. In general, AMPA receptors appear to be less affected by anesthetics than NMDA receptors. The exact mechanism of the anesthetic block of the NMDA receptor is unresolved but it has been shown that pentobarbital can block the open NMDA channel, which decreases the time the channel is open
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Table 15.4 Effects of General Anesthetics on Ligand-gated Channels Anesthetic
NMDA-receptor
AMPA-receptor
GABAA-receptor
Halothane
ø: [81,118, 322] I: [70,95,181] ø: [30a,96a] I: [70,180b,182,183,343]
ø: [118,322] I: [70,95,181] ø: [96a] I: [70,180b,182]
P: [94,205,324,340,342]
P: [215,217]
I: [197]
P: [187,215,217b]
I: [196,197,345]
I: [187] I: [182,186, 187] I: [69,70] I: [176–178] ø: [173,184,347]
ø: ø: ø: ø:
P: [68,182,187,202, 205,340,341,343, 344a,68a,94a,88a] P: [187] ø: [66] P: [187] ø: [182] P: [66,68] P: [205] ø: [218] P: [67,103,340] P: [67]
P: [187] P: [187] P: [67,217] ø: [217,218] P: [67,103,205,217] P: [67] ø: [217]
I: [187] I: [187] I: [195,346] I: [218,346] I: [196,197] ø: [346]
Isoflurane
Nitrous oxide Xenon Barbiturates Ketamine Propofol Etomidate
[182,186] [185] I: [70] [179] [185,347]
Glycine-receptor
nACh-receptor
Considered are channels that are involved in fast neurotransmission. Included are studies where receptors were expressed in non-neuronal cells. Only studies that use anesthetic concentrations in the clinical range, i.e., 52 MAC for volatile anesthetics and 52 ED50 for intravenous anesthetics are listed. Anesthetic effects are only considered relevant when they exceed a 10% change from baseline channel function (compare Franks, N.P. and Lieb, W.R., Toxicol. Lett., 100–101, 1–8, 1998.) (P: potentiation of receptor function; ø: no effect on receptor function; I: inhibition of receptor function). a Experiment conducted with sevoflurane. b Experiment conducted with enflurane.
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to allow an inward current [69]. Similarly, dissociative anesthetics like ketamine preferentially block open NMDA channels [177,178]. Some neuronal systems such as spinal cord motoneurons that strongly depend on glutamatergic sensory afferent inputs have been shown to be sensitive to anesthetic agents, and in fact the nocifensive withdrawal reflex that is used for MAC determination of volatile anesthetics is dependent on these glutamatergic inputs. Cheng and Kendig have convincingly demonstrated that various volatile anesthetics including enflurane cause a significant depression (30–35%) of both the NMDA and AMPA type of postsynaptic ionotropic glutamate receptor response at 1 MAC in these spinal motoneurons [180]. Lastly, nitrous oxide and xenon are two gaseous anesthetics, which both possess relatively weak amnesic/anesthetic actions, but exhibit clinically useful analgesic effects at subanesthetic concentrations. These agents have recently been shown to almost exclusively depress postsynaptic glutamate receptors with little effect on other ligand-gated channels [182,186,187]. For example, 1 MAC xenon (71%) decreased the total charge transfer through the excitatory postsynaptic current in autaptic cultures of hippocampal neurons by 60% by selectively inhibiting the NMDA-mediated component of the current [182]. Similarly, when 0.5 MAC concentrations of nitrous oxide (58%) and xenon (46%) were tested on a variety of recombinant brain receptors expressed in Xenopus oocytes, the NMDA receptors were the most sensitive and were inhibited by about 30–35% by both agents [187]. Unlike ketamine, the effect of gaseous anesthetics on NMDA receptors was not voltage-dependent, suggesting that their mechanism of action at the receptor level differs from that of ketamine. For a detailed review of anesthetic actions on excitatory amino acid receptors we refer the reader to Ref. [188]. Neuronal Nicotinic Acetylcholine Receptors
Neuronal nicotinic acetylcholine (nACh) receptors belong to a family of acetylcholine-gated, cation-selective channels consisting of combinations of homologous subunits to form a pentameric structure. It is noteworthy that nACh receptor subtypes containing the a4/b2 subunits are particularly prevalent in the CNS, but it is unclear whether they have a major function as postsynaptic receptors. In general, nACh receptors are not directly involved in fast excitatory neurotransmission between CNS neurons, but it has been suggested that they modulate the release of other neurotransmitters via a location on presynaptic nerve terminals [189]. While the exact role of neuronal nACh receptors for the anesthetic state remains unresolved it has been suggested that cholinergic receptors play a significant role in the respiratory system both peripherally and centrally [190]. Stimulation of nicotinic cholinergic receptors in the carotid body chemoreceptors [191]
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increases carotid sinus nerve discharge, and nicotine administered locally in the ventrolateral medulla, i.e., to the area of presumed central chemoreceptors, causes large increases in ventilation [192]. However, in contrast to these findings, Haji et al. found that feline laryngeal respiratory motoneurons, excited by iontophoretically-applied acetylcholine, could only be blocked with the muscarinic antagonist atropine but not the nicotinic antagonist hexamethonium [193]. In addition, it has been suggested that muscarinic cholinergic mechanisms may be more relevant to central chemosensing than nicotinic mechanisms [194]. Various intravenous [195], volatile [196,197] and gaseous [187] anesthetics inhibit acetylcholine-mediated channel activation by decreasing the time neuronal nicotinic channels are open, but the exact mechanism is unresolved. Two groups showed that brain nACh receptors of the a4/b2 subtype composition are particularly sensitive to subanesthetic concentrations of volatile anesthetics [196,197]. Flood et al. found the IC50 of isoflurane that inhibited the a4/b2 receptors expressed in Xenopus was 85 7 mM (i.e., 0.3 MAC) [196]. Violet et al. showed in a similar system (nicotinic muscle and brain receptors expressed in Xenopus oocytes) that halothane, isoflurane and sevoflurane depressed the neuronal a4/b2 subtype at subanesthetic concentrations (IC50 0.1–0.3 MAC) while propofol was relatively ineffective [197]. Similarly, the a4/b2 subtype of recombinant nACh receptors expressed in Xenopus oocytes was depressed 39% by 0.5 MAC nitrous oxide [187]. Recently, neuronal nACh receptors have been ruled out as playing any role in anestheticinduced immobility [198], since a4/b2 nACh receptors expressed in Xenopus oocytes are also potently inhibited by nonimmobilizing perhalogenated alkanes [199]. Inhibition of a4/b2 nACh receptors has been suggested to mediate amnesic action and suppress learning at subanesthetic concentrations [199]. Interestingly, nonhalogenated alkanes potently inhibit nACh receptors but do not affect the agonist dissociation constant of these receptors nor of the structurally related GABAA receptors [200]. GABAA Receptors
Among the ligand-gated receptors, the GABAA receptor plays a dominant role in fast inhibitory neurotransmission, and this holds true for the brainstem respiratory network [201] (see section IV.B). The prominent effect of all modern volatile anesthetics [68,202] and many intravenous anesthetics, in particular propofol [67,103], barbiturates [67,68], and etomidate [67], is a pronounced increase in GABAA receptor function by increasing the receptor affinity for GABA, and thus potentiating the effects of submaximal GABA concentrations. It is
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of note that some anesthetics can gate the GABAA channel directly in the absence of GABA and cause inhibitory postsynaptic currents [202]. Anesthetics prolong open channel times and increase the chloride conductance of the channel that is gated by the GABAA receptor [202], resulting in an increase in the negative charge transfer across the membrane of the postsynaptic neuron [203,204]. In Xenopus oocytes the enhancement of GABA-induced currents at low levels of GABA (5 mM) ranged from 150% for clinically relevant (clinical ED50) concentrations of isoflurane to 200% for halothane and over 300% for enflurane [205]. In all cases the total charge transfer was greatly increased by prolongation of the slow component of the inhibitory postsynaptic current, causing neuronal depression [182,202,205,206]. The neuronal depression resulted from a decrease of neuronal excitation that was either due to shunting of excitatory postsynaptic potentials (EPSPs) or to hyperpolarization of the neuronal resting membrane potential or both. The exact mechanisms of the intravenous and volatile anesthetic effects on GABAA receptors have been studied in considerable detail [66,182,202,206,207]. Interestingly, inhibitory postsynaptic potentials (IPSPs) mediated by synaptic GABAA receptors have a much faster decay time than those mediated by extrasynaptic receptors [208]. Brickley et al. suggest that extrasynaptic receptors are activated by ambient GABA that might have spilled over from synapses, causing tonic inhibition of the neuron, while synaptic receptors mediate fast transient inhibition [209]. This could result in differential anesthetic effects on the excitability of a neuron [208]. Lastly, some anesthetics, for example isoflurane, show a clear stereoselective effect at the GABAA receptor [210] as well as in the whole animal [211]. Such stereoselectivity of receptor effects and clinical actions suggests that the GABAA receptor is an important protein target for the anesthetic action of isoflurane [138]. Glycine Receptors
Glycine plays a dominant role as an inhibitory neurotransmitter in the spinal cord [212] while its effects at supraspinal sites are less well defined [213]. Schmid et al. have described effects of glycine in bulbar respiratory neurons [214]. The responses of recombinant glycine receptors to low concentrations of glycine, expressed in Xenopus oocytes, as well as native glycine receptors in rat medullary neurons, were potentiated by clinical concentrations of volatile anesthetics [215]. Sevoflurane was more potent than halothane, isoflurane, and enflurane. The potentiation of glycinergic currents at 1 MAC was 20–80%, less than that seen for GABA in similar systems, but the overall mechanism, i.e., an increase in charge transfer via prolongation of channel opening by the anesthetics, appeared similar
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[215–217]. Intravenous anesthetics such as propofol [103] and barbiturates [217], but not ketamine [217,218] or etomidate [217], also potentiate glycine receptor function. As in other ligand-gated receptors, anesthetic effects on glycine receptors are highly dependent on receptor subunit composition. Spinal glycine receptors containing the a1 subunit are relatively sensitive to propofol [103] while hippocampal glycine receptors containing the a3 subunit are quite insensitive [219]. C. Summary and Conclusions
General anesthetics may exert their effects via an almost unlimited number of potential neuronal targets. However, if sensitivity to clinically relevant concentrations is considered, a superfamily of ligand-gated ionotropic receptors stands out that is important for fast neurotransmission at central synapses. The superfamily of genetically related receptor channels comprises the excitatory nicotinic acetylcholine receptor and the inhibitory GABAA and glycine receptors and is by far the most anesthetic-sensitive. GABAAergic inhibition plays a universal role in the CNS while nicotinic and/or muscarinic cholinergic neurotransmission seems to play a role in sedation/unconsciousness [220], learning [220], and chemosensitivity [15]. The anesthetic potentiation of GABAA receptors is powerful, and neuronal nicotinic acetylcholine receptors show supersensitivity to anesthetic depression [221]. Ionotropic glutamate receptors do not belong to the above superfamily. However, glutamate receptors are very important for excitatory neurotransmission in the CNS and the brainstem respiratory network (see section IV.B). The AMPA subtype of the glutamate receptor seems to be relatively insensitive to most anesthetics while the NMDA subtype of glutamate receptors is strongly affected by a large group of anesthetics. This receptor subtype has been implicated in learning, nociception, and chemosensitivity. Voltage-gated cation channels, including Naþ, Kþ, and most subtypes of Caþþ channels, tend to be relatively insensitive neuronal targets. However, a depression of sodium and calcium channels may play a role in the anesthetic reduction of presynaptic transmitter release. The recent discovery of background tandem pore Kþ channels that are activated by volatile anesthetics at clinical concentrations and the prediction that these channels may be abundant in the CNS suggests that these channels could be highly relevant for explaining general anesthetic effects. Such channels have been recently described in HMs. The relevance of this model to anesthetic effects on respiration will be discussed in further detail in section V.C.
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Overview of the Brainstem Respiratory Network
A. Current Model of the Adult Mammalian Respiratory Network
Automatic ventilation in contrast to voluntary ventilation is regulated primarily by mechanisms in the medulla and pons. These areas contain respiratory neurons responsible for the generation of rhythmic discharge patterns that are relayed to respiratory motoneurons, which finally innervate the respiratory musculature [88,222–229]. Respiratory motoneurons include spinal motoneurons, which innervate muscles of the respiratory pump (diaphragm, intercostals, and abdominal muscles), and cranial motoneurons, which innervate muscles that modulate airway resistance [230]. The latter include laryngeal and pharyngeal motoneurons with somata in the nucleus ambiguus and axons in the vagus nerves and HMs, which innervate the striated muscles of the tongue. Respiratory-related neurons in the medulla are concentrated in two regions (Figure 15.6). The DRG is localized ventrolateral to the nucleus solitarius, and the VRG is localized near the nucleus ambiguus. The great
PRG DRG
nA
Böt. C. pre-Böt. C.
nA Böt. C.
rostral VRG
RVLM IX, X
Obex
caudal VRG (nRA)
nTS
DRG nA rostral VRG
Phrenic Int. Intercostal
Ext. intercostal
Figure 15.6 Brainstem and spinal cord locations of the major groups of respiratory-related neurons. Left: transverse sections of the medulla at the levels indicated by dashed horizontal lines superimposed on a dorsal view of medulla (right). Bo¨t. C.: Bo¨tzinger complex; pre-Bo¨t. C.: pre-Bo¨tzinger complex; PRG: pontine respiratory group; DRG: dorsal respiratory group; VRG: ventral respiratory group; nA: nucleus ambiguus; nRA: nucleus retroambigualis; nTS: nucleus tractus solitarius; Int.: internal; Ext: external; IX, X: 9th and 10th cranial nerves; RVLM: rostral ventrolateral medulla (adapted from Ref. 244).
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majority of the DRG neurons are bulbospinal I premotor neurons, which innervate phrenic and intercostal motoneurons [226]. The VRG contains bulbospinal neurons, glossopharyngeal and vagal motoneurons, and neurons with axonal projections that remain within the brainstem, termed propriobulbar. The caudal part of the VRG is mainly composed of expiratory (E) premotor neurons located in the nucleus retroambiguus. Premotor neurons play a key role in pattern generation (see below). They relay phasic excitatory drive, either monosynaptically or polysynaptically, to their respective motoneurons [231–234] and also appear to produce inhibition of antagonistic motoneurons via interneurons [235] to prevent simultaneous activation of inspiratory and expiratory muscles during eupnea. In its rostral part, the VRG contains mostly I neurons and extends further rostrally to the Bo¨tzinger complex, a group of mainly E neurons located just dorsal of the facial nucleus. A region immediately caudal to the Bo¨tzinger complex that contains a mixture of propriobulbar neurons with inspiratory, expiratory, and phase-spanning activities is known as the preBo¨tzinger complex. In vitro studies in neonatal preparations suggest that the preBo¨tzinger complex contains the neurons necessary for the generation of respiratory rhythm. Using a series of precisely controlled neuraxis transections of an isolated brainstem-spinal cord preparation from neonatal rats, Smith et al. [236] identified a region (preBo¨tzinger) whose destruction led to the elimination of rhythmic motor output. To rule out the possibility that the transections separated a neuronal network essential for rhythm generation, they demonstrated that rhythm was still present in transverse brain slices containing the preBo¨tzinger area. They also found neurons in the ventral part of such slices that discharged in synchrony with the burst activity of the hypoglossal nerve roots and some neurons whose rhythm could be altered by manipulation of their membrane potential, concluding that these neurons constituted pacemakers. These results have been confirmed by several more recent studies [87,236–240]. In contrast to the pacemaker theory, some authors suggest that the preBo¨tzinger complex contains a network of reciprocal inhibitory neurons that translates chemodrive into respiratory rhythm [241–245] (see below). It is possible that medullary rhythm generation changes during ontogenesis from a pacemaker-driven system at the fetal and neonatal stage to a network requiring postsynaptic inhibition in the mature brain [229,245]. Additional respiratory-related neurons are found in high concentration in the pons, next to the nucleus parabrachialis medialis and the Ko¨lliker-Fuse nucleus. This rostral pontine region has been called the pneumotaxic center or the pontine respiratory group. It provides additional drive to the medullary respiratory group and may support rhythm generation by enhancing phase switching between the inspiratory and expiratory phases [225].
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Drive and Timing of the Respiratory Motor Pattern
From a functional point of view, respiratory output is defined by two components: respiratory drive and respiratory rhythm or timing. Recording from the purely inspiratory phrenic nerve can assess these components. The peak phrenic amplitude represents the respiratory drive and is a good estimate of diaphragmatic inspiratory force [31], while the timing is reflected in the number and duration of inspirations per minute. The main sources of drive in automatic ventilation are the central and peripheral chemoreceptors. The effect of increasing central CO2 drive via rebreathing during hyperoxia in the vagotomized dog results in an increase in peak phrenic activity and peak transpulmonary pressure without any change in the respiratory cycle duration (unpublished observation, E.J. Zuperku, 1986, Figure 15.7A). In contrast, in vagally intact dogs, changes in intrathoracic pressure activate slowly adapting PSRs located in airway smooth muscle, which can produce profound changes in I and E phase durations (TI and TE) with little effect on drive. For example, an increase in positive end-expiratory pressure can prolong the E phase without effect on the peak phrenic activity or I phase duration (Hering-Breuer expiratory facilitatory reflex, Figure 15.7B1). An increase in PSR activity during the I phase produces a shortening of the I phase. This results in a reduction of peak phrenic activity although the drive is not changed (Hering-Breuer inspiratory inhibitory reflex, Figure 15.7B2). In addition, the reflex shortening of the I phase produces a shortening of the subsequent E phase via a central mechanism [247,248]. Activation of peripheral chemoreceptor afferents in the canine aortic depressor nerve (cranial nerve X) produces a profound reduction in both I and E durations without change in peak phrenic activity (Figure 15.7C) [249]. Stimulation of the carotid body chemoreceptor (cranial nerve IX) via a CO2-saturated saline bolus injected into the common carotid artery results in a brisk increase in peak phrenic activity together with an increase in inspiratory duration and a shortening of the subsequent expiratory phase (Figure 15.7D). Interestingly, in this last example peripheral chemoreceptor activation affected both drive and timing. Tonic Respiratory Drive
A minimum CO2 drive sensed by central and peripheral chemoreceptors [250] is necessary to maintain normal interactions among the different types of respiratory neurons [15]. Iontophoresis of tetrodotoxin (TTX), which interrupts synaptic transmission by blocking Naþ channels, eliminates neuronal discharge and causes a constant hyperpolarization in virtually all types of respiratory neurons [251,252]. This suggests that phasic discharge is principally due to synaptic interactions among respiratory neurons.
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PNG 30 sec
0 Pt (cm H2O)
Pt (cm H2O)
-5 -10 -15
B1
10
B2
5 20
40 100
5 sec
0 Hz
0
PNG 1 sec
C
ADN stimulation (5 Hz)
D
PNG 1 min
co2/saline bolus
5 sec
Figure 15.7 Drive and timing components of the breathing pattern illustrated by the phrenic neurogram (PNG) responses to various respiratory-related inputs. (A) Drive aspect shown by the effect of increasing central CO2 drive via rebreathing during hyperoxia in the vagotomized dog. Pt: transpulmonary pressure. (B1) Timing aspect in vagally intact dogs, where an increase in positive end-expiratory pressure increases slowly adapting PSR activity, which reflexly increases the duration of the E phase (TE) without affecting the peak phrenic activity or I phase duration (TI). (B2) An increase in electrically induced (Hz) PSR activity during the I phase reduces TI and as a result reduces peak phrenic activity. (C) Timing effects only. Electrically induced aortic chemoreceptor activity shortens both I and E phase durations with negligible effect on peak PNG. (D) Combined drive and timing effects. Stimulation of the carotid body chemoreceptors via a CO2 saturated saline bolus injected into the common carotid artery increases peak phrenic activity together with an increase in inspiratory duration and a shortening of the subsequent expiratory phase.
Tonic excitatory inputs maintain the membrane potential between 50 and 70 mV, which allows additional excitatory inputs to depolarize the neuron above firing threshold [170]. Hypercapnia stimulates virtually all brainstem respiratory neurons [253,254]. Normally, ventilation doubles for a 2–4 mm Hg rise in PaCO2 [15,255]. In anesthetized mammals including man, hypocapnia through artificial hyperventilation can result in apnea where inspiratory neurons become quiescent and expiratory neurons fire tonically [256–258]. However, during wakefulness hypocapnia does not lead to sustained apnea, which suggests a wakefulness- or state-dependent drive that is derived from the reticular formation [259,260].
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Peripheral chemoreceptors, located in the carotid and aortic bodies, supply about 30% of the CO2-related drive for ventilation under normoxic conditions [261,262]. The greater amount of CO2 drive arises from central chemoreceptors. Traditionally their main location is believed to be on the ventral medullary surface since direct application of acidic artificial CSF to this area stimulates breathing [263,264]. However, the more recent view is that central chemoreceptors are more widespread within the medulla [15]. In vivo studies, employing focal microinjections of the carbonic anhydrase inhibitor acetazolamide, which produces a localized acidosis, and microdiffusion of CO2, are able to locate brainstem regions that result in reversible increases of phrenic nerve activity. These regions include sites along the ventral medulla within 800 mm of the surface, near the NTS, near the locus coeruleus, the medullary raphe´, and the VRG region [15]. A Functional Model for Rhythm Generation in the Adult Mammal
The variety of neuronal discharge patterns and the precise control of the timing of I and E phases in adult in vivo preparations have led to the development of neural network models where rhythm generation is based on inhibitory synaptic interactions [225,226,229,244]. This hypothesis is supported by the observation that breathing rhythm in mature rats is interrupted when GABAergic and glycinergic synaptic currents are reduced by a reduction of chloride ions in the arterial perfusate of an in situ brainstem-spinal cord preparation [265]. Similarly, bath application of strychnine, a blocker of the inhibitory glycine receptor, caused a disruption of rhythm in medullary slices from adult rats [246]. In contrast, in the isolated brainstem-spinal cord preparation from neonatal rats, block of the chloride-dependent postsynaptic inhibition modulates but does not halt rhythm [246,266]. This points to developmental changes in rhythm generation [229,245]. Reflex control of discharge patterns is also mediated at the level of the premotor neurons. For example, E bulbospinal neurons are highly responsive to central and peripheral chemoreceptor stimulation [267,268]. In addition, these E premotor neurons receive inputs from slowly adapting PSRs. They are excited by lung inflation in the low volume (or transpulmonary pressure) range but become progressively inhibited with further increases in lung volume [269,270]. The discharge pattern of E premotor neurons is mainly determined by the time-course pattern of lung volume during the E phase, and their bidirectional response to the pulmonary afferent input may play a role in the control of expiratory airflow rate and end-expiratory lung volume [269]. All these observations have led to the development of a functional model for the transformation of mainly tonic drive into appropriate motor discharge patterns (see Figure 15.8). Tonic excitatory drive is relayed to all
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I Premotor
Central chemosensors Peripheral chemoreceptors State-dependent & other
I Motor
I pattern shaping/ control
Excitatory Drive mainly tonic
SYNAPTIC INPUTS Excitatory Inhibitory (+/-) bidirectional Opposite phase inhibition AFFERENT INPUTS
E timing
I timing
Rhythmic Discharge Patterns
E pattern shaping/ control
E Premotor
E Motor
Excitatory and/or Inhibitory (Modulating inputs not shown)
Figure 15.8 Functional diagram of the ponto-medullary respiratory pattern generator that transforms tonic excitatory drive into rhythmic neuronal discharge patterns. See text for further details.
elements of the network to bring them to their operating levels. The drive that reaches the motoneurons is relayed via premotor neurons. At the level of the premotor neurons the tonic drive is interrupted by strong periodic inhibition from the phase-timing component of the network. This phasetiming component (E timing/I timing, Figure 15.8) is believed to be located in the preBo¨tzinger complex. Figure 15.9A presents a model for a neuronal network oscillator, a simplified version of the model originally proposed by our laboratory [241] and other more recent models for respiratory rhythm generation [242–245]. The model is based on reciprocal inhibition between pools of I and E decrementing neurons (Figure 15.9B) and modeled by a chord-conductance representation of a neuronal membrane (Figure 15.9C). The membrane includes conductances for synaptic inhibition (Gsyn), e.g., GABAA receptors; excitatory drive (Gexc), e.g., glutamate receptors; pooled leak channels (Gleak), e.g., leak Cl, Naþ and Kþ channels; and intrinsic membrane stabilization via voltage-dependent channels with slow dynamics (Gvolt), e.g., a slow, delayed, outward rectifier. The latter produces a postinhibitory rebound effect, which results in an adapting (i.e., decrementing) pattern of inhibition that is eventually overwhelmed, allowing the phase to switch. The behavior of this model can mimic many of the salient functional characteristics of the hypothetical brainstem oscillator, i.e., decrementing neuronal discharge pattern and phase timing.
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In section IV.C, we will describe how the effects of anesthetics on the membrane conductances alter respiratory timing in the oscillator in a range that is also observed in vivo. The oscillator provides synaptic outputs that completely inhibit the discharge of the premotor neurons during the silent phase (the E phase of I premotor neurons and the I phase of E premotor neurons). Additional neuronal circuitry is required to shape and control the final patterns, i.e., augmenting and decrementing, of the premotor neurons. Due to the strong correlation between the slopes of these patterns and respiratory phase timing, it is assumed that a bidirectional interaction between the timing and pattern-shaping components must exist (Figure 15.8). The following section will discuss the roles of different receptor subtypes for specific functions in respiratory timing and pattern generation. B. Neurotransmitters Involved in Rhythm and Pattern Generation Glutamate
Excitatory neurotransmission within the respiratory network is essentially mediated by glutamate, while inhibitory neurotransmission is mediated by GABA and glycine [170]. Since ionotropic glutamate, glycine, and GABA
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receptors are found on almost all respiratory neurons, systemic application of their agonists and antagonists can elicit profound changes in breathing pattern, but determination of their specific roles is impossible. Thus, our discussion will be restricted to studies using highly localized methods of application, such as microiontophoresis and pressure microejection. Respiratory neurons typically show a specific augmenting, decrementing or constant discharge pattern that depends on the neuronal membrane characteristics and synaptic inputs (e.g., Figure 15.10). In all types of respiratory neurons, EPSPs are present during the active phase of their
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respiratory cycle [227,271–276]. Glutamate or selective NMDA and AMPA agonists increase action potential firing rate in these neurons [47,81,95, 277–284]. Iontophoresis of 2,3-dihydroxy-6-nitro-7-sulphamoylbenzo (F) quinoxaline (NBQX), an AMPA receptor antagonist, or dizocilpine, an NMDA receptor antagonist, decreases the active phase depolarization together with an increase in input resistance and suppresses action potential firing [170]. NMDA agonists appear to be less effective during the inactive phase of the neuron than AMPA agonists, which may be due to the voltage-dependent properties of the NMDA receptor [170]. This suggests that sequential activation of postsynaptic AMPA and NMDA receptors is required for full neuronal discharge activity. However, studies from our laboratory suggest that AMPA-receptor activation is not required for physiological function of canine bulbospinal E neurons [280,282,285]. Picoejection of the AMPA receptor antagonist NBQX on bulbospinal E neurons has no effect, while picoejection of the NMDA receptor antagonist AP5 reduces the discharge activity of these neurons to near zero. These results suggest that AMPA receptors on bulbospinal E neurons are not endogenously active. Similar studies show that on bulbospinal I neurons, each glutamate receptor subtype plays a distinctly different role: tonic drive is exclusively mediated by NMDA receptors while phasic excitatory drive is exclusively mediated by AMPA receptors [282,285]. The NMDA receptor-mediated tonic drive contributes about two-thirds of the total drive, while AMPA receptor-mediated phasic drive contributes the other one-third (see Figure 15.11). The latter is thought to be due to self-reexcitation, i.e., the reciprocal excitatory coupling among augmenting I neurons during the inspiratory phase. It has been suggested that NMDA receptors are involved in the change from inspiration to expiration, i.e., the inspiratory off-switch, because systemic injection of the NMDA receptor antagonist dizocilpine (MK-801) results in apneusis in vagotomized cats [50,65,286]. Since animals with intact vagi, which relay PSR input directly to the medullary respiratory center, show no change in respiration, the affected mechanisms are attributed to an inspiratory-off-switch-related pontine structure [286]. Accordingly, microinjection of NMDA antagonists into the pontine area induces apneusis [287]. However, microinjection of NMDA antagonists into the nucleus tractus solitarius (NTS) also evokes apneusis [288,289]. In addition, pneumotaxic-disconnected and vagotomized cats can maintain eupneic respiration but show apneusis after intravenous dizocilpine [290]. Taken together, NMDA receptor mechanisms play an important role in inspiratory off-switch, not only in the pontine structure, but also in the medullary respiratory network. Metabotropic glutamate receptor activation, which modulates synaptic transmission through G-protein-coupled signal transduction, has no significant effect on the membrane potential of bulbar respiratory neurons.
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Figure 15.11 Neuronal responses of a single inspiratory neuron in vivo to neurotransmitter antagonists given in the order: (1) picrotoxin, (2) NBQX, (3) AP5. (A) Picrotoxin-induced activity during the expiratory phase with no effect during the inspiratory phase. (B) NBQX reduced inspiratory phase activity but not the tonic expiratory phase activity. (C) AP5 reduced the activity in both phases (adapted from Ref. 285).
Rather, it appears that the transmission of inspiratory drive is fine-tuned by metabotropic glutamatergic modulation at the level of the spinal cord [170]. GABA and Glycine
In all main types of respiratory neurons, IPSPs can be recorded during the inactive phase of the respiratory cycle. Decrementing and augmenting I neurons receive IPSPs during expiration; the subgroup of late-I neurons also receives IPSPs during early inspiration. Decrementing E neurons receive IPSPs during inspiration and expiration and augmenting E neurons during inspiration and early expiration [170]. GABA mediates IPSPs during the inactive phase through GABAA receptors. These results are in agreement with extracellular recordings where GABAA and glycine antagonists were applied to single respiratory neurons [214,291,292]. Active phase IPSPs have been demonstrated in augmenting I and augmenting E neurons. These IPSPs are identified as a decreased input resistance and reversal of membrane potential near the equilibrium potential of Cl, indicating the involvement of GABA or glycine receptors [97,98,251,293]. Extracellular studies in vagotomized cats [214] suggest that the inhibition during the respiratory phase switch in augmenting I and
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augmenting E neurons is mediated by glycine, since the iontophoresed glycine antagonist strychnine induces action potentials selectively at that time while the GABAA antagonist bicuculline has little or no selective effect [291]. It has also been suggested that glycine is the major source of inhibition during the early expiratory phase in most I and E neurons [214]. However, the IPSPs in augmenting I neurons during inspiratory offswitch evoked by vagal stimulation are increased by systemic midazolam, a benzodiazepine GABAA modulator, and decreased by systemic bicuculline but not strychnine [65,294]. Similarly, the inspiratory off-switch is facilitated by midazolam and impaired by bicuculline. GABAAergic inhibition that can be blocked with picrotoxin is also responsible for silent phase inhibition in canine I and E bulbospinal neurons. Picoejection of strychnine produces little or no effect [285,292]. In addition to silent phasic inhibition, in some neuron types, extracellular iontophoresis of bicuculline or strychnine depolarizes the membrane throughout the whole respiratory cycle [252]. These effects are not seen after iontophoresis of TTX, suggesting that the tonic inhibition is mediated by GABAA and glycine receptors and may serve to preserve the neuronal membrane potential level. Another key role for tonic GABAA receptormediated inhibition is gain modulation of neuronal discharge patterns [292,295,296]. This mechanism constrains the control- and reflexly-induced activities of respiratory neurons to about 35–50% of the discharge rate without inhibitory input. GABAergic gain modulation may provide a mechanism for the optimal control of respiratory as well as other behavioral functions (e.g., coughing, sneezing, vomiting) mediated by respiratory premotor neurons [295]. In summary, both GABA and glycine are essential for generating respiratory rhythm in the mature respiratory network. Additional Neuromodulation in the Respiratory Network
GABA also activates bicuculline-insensitive GABAB receptors, which increase postsynaptic Kþ conductances or decrease presynaptic Ca2þ conductances [297–299]. Iontophoresis of the GABAB receptor agonist baclofen hyperpolarizes the membrane during the whole respiratory cycle, with a decrease in input resistance and spiking [300–302]. Iontophoresed antagonists by themselves increase the spike activity of respiratory neurons [302], suggesting that GABAB receptor-mediated tonic inhibition is present in respiratory neurons. A presynaptic contribution of GABAB receptors is suggested for phrenic motoneurons [230]. The activity of motoneurons is further controlled by sensory inputs and synaptically mediated neuromodulators. The synaptic control of phrenic motoneurons is complex, and they are not simple relays for respiratory control of the diaphragm. There is evidence for postsynaptic receptors for serotonin (5-HT), norepinephrine, neurokinins, neuropeptide Y,
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galanin, metenkephalin, cholecystokinin (CCK), and thyrotropin-releasing hormone (TRH) and for presynaptic receptors for 5-HT, GABAB, metabotropic glutamate and adenosine [298,299]. Hypoglossal, laryngeal, and other respiratory-related cranial motoneurons also contain receptors for many of the same neuromodulators [298]. While the specific function of each neuromodulator is unknown, it has been suggested that their presence could provide a means for controlling neuronal excitability over a broad time scale, from tens of milliseconds to years [299]. They may have a role in adaptation to physiological state, developmental, environmental, pathological, and mechanical changes that require a sustained change in respiratory function. C. Possible Mechanisms for Anesthetic-induced Changes in Rhythm Oscillator Model for Central Anesthetic Effects on Respiratory Rate
Inhalational anesthetics cause an increase in breathing rate [21,72–76], while some IV anesthetics such as barbiturates and propofol can cause a slowing in respiratory rate in rabbits and cats, although not in humans [71,83,84]. Possible sites of anesthetic action include inputs from sensory afferents (e.g., PSRs), CNS inputs from outside the medulla, and direct effects on intramedullary rhythm-generating mechanisms. In the decerebrate, vagotomized dog model where suprapontine inputs and pulmonary afferents are removed, halothane and sevoflurane still markedly increase breathing rate (e.g., Figure 15.19 and Figure 15.20 [30,81]). This suggests that the anesthetic effects on respiratory rhythm take place within the pontine-medullary rhythm-generating network. Simulation studies of the network oscillator model presented in section IV.A demonstrate that anesthetic effects on the rhythm-generating network alone could indeed result in similar respiratory frequency changes as those observed in vivo [30,81,94]. Reduction of tonic excitatory drive by 20% causes an increase in respiratory frequency while peak neuronal discharge frequency is reduced (Figure 15.12A). The rate effect resembles the tachypnea that can be observed with a reduction in CO2 chemodrive (see also section II.B). In contrast, a 20% increase in reciprocal synaptic inhibition to the oscillator results in a decrease in respiratory frequency while the neuronal discharge frequency is increased (Figure 15.12B). Studies on neurotransmission in respiratory premotor neurons suggest that volatile anesthetics reduce overall excitatory drive to the neurons while overall synaptic inhibition is increased [30,81,86,88,95,96]. In our computer model, the combination of a 20% decrease in excitatory drive with a 20% increase in reciprocal synaptic inhibition, which is in the range of the effects observed in premotor neurons, leads to an increase in respiratory frequency together with a decrease in neuronal discharge frequency (Figure 15.12C). These effects are consistent
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Figure 15.12 Predicted changes in respiratory rhythm when the magnitude of synaptic inputs to the simulated central oscillator (Figure 15.9) is altered. (A) Effect due to changes in tonic excitation. (B) Effect due to changes in reciprocal inhibition. (C) Effect due to combined changes in tonic excitation and reciprocal inhibition. See text for details.
with in vivo results. Thus, it is possible that some of the anesthetic effects on respiratory rhythm result from modest changes in chemodrive and synaptic transmission within the medullary rhythm-generating network. Rate Changes Produced by Anesthetic Effects on Pulmonary Afferents
Changes in pulmonary receptor activity may also contribute to the tachypnea observed with inhalational anesthetics. Dogs were placed on cardiopulmonary bypass so that the lungs were isolated from the systemic
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Figure 15.13 Direct effects of halothane on pulmonary receptors and resultant responses in the respiratory rate. (A) Tachypneic response to halothane insufflation of the circulatory-isolated lungs of a dog on cardiopulmonary bypass. Upper trace: airway halothane concentration; middle trace: transpulmonary pressure; lower trace: diaphragm EMG. (B) Effect of halothane on the PSR activity of a few-fiber preparation. See text for details.
circulation and thus from the systemically applied anesthetic, which allowed independent pulmonary administration of different concentrations of halothane. Increasing intrapulmonary halothane concentrations produced tachypnea in 17 of 21 dogs (see Figure 15.13A) [303]. The onset of the response is very rapid and the recovery phase is also short. Vagotomy eliminated the responses to halothane insufflation. Studies of the effects of halothane on the response characteristics of the slowly adapting PSRs to controlled lung inflations, suggest that these effects, acting via the Hering-Breuer reflex, produce the tachypnea described above. The relationship between PSR discharge frequency and transpulmonary pressure (Pt) consists of an activation pressure threshold below which the fiber is silent, a linear range above the activation threshold, and at higher pressures a saturation range in which the PSR has reached its maximum discharge rate [304,305]. The PSRs that control TI and TE appear to be located in the more distal airways [306], even though there is a large of number receptors in the larger conducting airways [304,307]. About 64% of the PSRs located in the canine distal airways have activation pressures near a Pt value equivalent to that at FRC (Pt ¼ 5–5.5 cm H2O) and 80% are activated at a Pt of 8 cm H2O [304]. In the absence of phasic inflations, small increases in FRC markedly increase TE without any effect
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on TI [308]. Thus, small changes in the PSR characteristics can affect breathing rate. The effects of halothane on the PSR characteristics have been studied in experiments using single- or few-fiber PSR activity, recorded from fine neural filaments of the distal portion of the transected cervical vagus nerve in anesthetized, paralyzed, and ventilated dogs [309]. An example of the effects of a progressive increase in the end-expiratory concentration of halothane from 0.5% to 3.0% on the discharge frequency of a PSR is shown in Figure 15.13B. Increasing halothane concentration increased the activation pressure (note the decrease of activity during the E phase), increased the sensitivity (F/Pt), and had no effect on the maximum discharge rate. These effects can be better seen in the discharge frequency (Fpsr) vs. transpulmonary pressure (Pt) plot of Figure 15.14, upper left, where each plot is based on 10 inflation cycles. Increasing concentrations of halothane shifted the activation threshold to higher pressures and increased PSR sensitivity (slope) in a dose-dependent manner. The pooled data (not shown) confirm these observations. Sensitization has also been observed for other agents such as enflurane, methoxyflurane, ether, chloroform, and trichloroethylene [309,310]. An increase in PSR sensitivity to lung inflation produces a shortening of TI via the volume threshold/ inspiratory off-switch mechanism described by Clark and von Euler [247] as depicted in Figure 15.14, upper right by the I function. As time progresses into inspiration, the declining timing function crosses a threshold (thr) upon which inspiration is terminated. During inflation, PSR activity increases this threshold, shortening TI from TI0 to TI1. An increase in PSR sensitization shortens TI to TI2. Associated with the shortening of TI is a shortening of TE via a central mechanism depicted in Figure 15.14, right, lower [247,248]. Thus, a reduction in TI and TE produces an increase in breathing rate ( f ). An additional potent mechanism controlling TE involves the level of PSR activity during and especially near the end of the E phase [311,312]. The anesthetic-induced increase in the activation pressure threshold will affect this mechanism by lowering PSR activity at end expiration. This effect on TE is illustrated by the E timing function (E ) of Figure 15.14, left, lower. Due to the nonlinear nature of the E timing function, small changes in threshold level produce marked changes in TE. Thus, increases in both PSR sensitivity and activation threshold can produce tachypnea via the Hering-Breuer reflex. Interestingly, increases in intrapulmonary CO2 concentration, which reduce PSR activity, also produce a rapid tachypneic response similar to that shown in Figure 15.13A [313]. A reduction in intrapulmonary CO2 concentration produces slowing. An effect of volatile anesthetics on other types of pulmonary receptors with vagal nerve fibers may contribute to tachypnea as well, in particular bronchial C-fibers [79]. It must be noted, however,
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Figure 15.14 Hypothesized mechanisms for changes in respiratory timing due to anesthetic-induced changes in PSR discharge characteristics. Upper left: Relationship between PSR discharge frequency (Fpsr) and transpulmonary pressure (Pt) for the PSR activities shown in Figure 15.13B. Halothane dose-dependently increases both PSR sensitivity (slope) and activation pressure (x-intercept). Right upper: mechanism for the Hering-Breuer I inhibitory reflex control of TI. I: central timing function, which determines TI when the threshold is crossed, also referred to as the volume-threshold function. Volume feedback via the PSRs increases the threshold (from thr), reducing inspiratory duration from TI0 to TI1. Halothane sensitization of the PSRs further increases the threshold, shortening inspiratory duration to TI2. TE is also reduced in proportion to TI due to an additional central mechanism (lower right). Lower left: mechanism for the Hering-Breuer E facilitatory reflex control of TE. E: central timing function, which terminates TE when the threshold (thr) is crossed. PSR activity lowers the threshold (*) and increases expiratory duration from TE0 to TE1. As indicated by the increase in PSR activation pressure, halothane reduces PSR activity (**) at FRC and thereby shortens expiratory duration from TE1 to TE2. Thus, both PSR sensitization and increased activation pressure produce a tachypneic response. The I and E decrementing discharge patterns (Figure 15.9, lower left) may serve as neural substrates for the I and E timing functions.
that the importance of the Hering-Breuer reflex in adult humans is still under discussion [84,220,314], although it seems relevant in infants [315], and that the tachypneic response to anesthesia can occur independently of a measurable Hering-Breuer reflex [71,75,316].
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Paradigms of Anesthetic Effects on Respiratory Neurotransmission
A. Respiratory Premotor Neurons: A Model to Explore Volatile Anesthetic Effects on Synaptic Neurotransmission In vivo
Zuperku et al. used an in vivo dog model to characterize neurotransmission to premotor neurons in the caudal VRG and the effects that various respiratory reflex inputs had on this neurotransmission [28–30,81,93,94,268,280,282,285,292,295,296,317–319]. Antidromic stimulation techniques showed that 90% of brainstem respiratory neurons whose soma was located approximately 1.5–3 mm caudal from the obex and 2.5–4.5 mm lateral from the midline projected axons to the spinal cord, i.e., they were respiratory premotor neurons [29]. The details of the preparation have been described elsewhere [28]. Studies on the effects of volatile anesthetics were conducted in decerebrate animals [320], which allowed the comparison of neurotransmission at 1 MAC of a volatile anesthetic with an anesthesia-free baseline state. These experiments were performed under steady-state conditions for respiratory chemodrive, i.e., hyperoxia and moderate hypercapnia (PaCO2 : 55–65 mm Hg). Neurotransmission to Respiratory Premotor Neurons
Premotor neurons are identified as inspiratory neurons when their discharge activity occurs simultaneously with the phrenic nerve activity and as expiratory when the phrenic nerve is silent. Application of the glutamate receptor agonists AMPA and NMDA increases the activity of both neuron types indicating that AMPA and NMDA receptors are present on the surface of the cell membrane [280]. Krolo et al. showed that about twothirds of the excitatory synaptic drive to inspiratory neurons is due to tonic excitation that is mediated by NMDA receptors [285] (Figure 15.15). The other third is due to phasic excitation and mediated by AMPA receptors [285]. This phasic drive appears to result from ‘self-reexcitation,’ i.e., inspiratory premotor neurons with augmenting discharge pattern reciprocally excite each other during the inspiratory phase [224]. In contrast, the excitatory synaptic drive to expiratory premotor neurons appears to be mediated solely by NMDA receptors [280]. In addition to the excitatory drive, respiratory premotor neuronal activity is determined by a tonic inhibitory input [295] and silent phase inhibition [292]. During the silent phase, i.e., the expiratory phase for inspiratory neurons and the inspiratory phase for expiratory neurons, neuronal activity is completely inhibited, mainly by GABAAergic [292] and to a small degree by glycinergic inhibition [285]. This phasic inhibition can be antagonized with the GABAA antagonist picrotoxin and the glycine antagonist strychnine [285] (Figure 15.16).
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Figure 15.15 Scheme of synaptic inputs to inspiratory and expiratory premotor neurons in the caudal VRG. Inspiratory bulbospinal neurons (IBSN) receive excitatory drive (Fe) via NMDA and AMPA receptors while expiratory bulbospinal neurons (EBSN) receive excitation only via NMDA receptors. The excitatory drive is gain modulated by bicuculline-sensitive GABAA-receptor-mediated tonic inhibition. The silent phase inhibition is mediated by picrotoxin-sensitive GABAA receptors. The neuronal discharge activity Fn during the active phase is determined by Fe and by the degree of GABAergic inhibition.
In addition to the silent phase inhibition, respiratory neurons are tonically inhibited via GABAA receptors that are only antagonized by the GABAA antagonist bicuculline [292] (Figure 15.15). This tonic GABAergic input attenuates the neuronal firing pattern such that antagonism of the inhibition with bicuculline increased the neuronal discharge frequency proportionally
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Figure 15.16 Diagram of the hypothetical components of the typical step-ramp discharge pattern of inspiratory premotor neurons in the caudal VRG. The components can be isolated by the application of the respective antagonists. Block of the silent phase inhibition with picrotoxin reveals the tonic drive to the neuron. The additional phasic excitation can be blocked with the AMPA receptor antagonist NBQX. Block of the tonic drive with the NDMA antagonist AP5 nearly stops the neuronal discharge (adapted from Ref. 285).
to the control discharge pattern [292]. The underlying phenomenon is known as gain modulation (Figure 15.17). Further investigation suggests that the GABAA receptors that mediate tonic inhibition are located closer to the soma than the AMPA and NMDA receptors that mediate the physiologic excitatory drive [296]. There are, however, additional AMPA and NMDA receptors located on or near the soma, since picoejection of small amounts of these glutamate agonists produces a rapid and marked neuronal excitation [321] (Figure 15.18). Effects of Volatile Anesthetics on Respiratory Neurotransmission
The effects of anesthetics on excitatory and inhibitory synaptic transmission have been studied with the picoejection technique. With the picoejection method, receptor agonists and antagonists can be ejected onto single neurons in vivo while recording their discharge activity extracellularly, i.e., the method allows measurement of the overall excitatory and inhibitory drive to a single neuron and its postsynaptic receptor response. Effects on the presynaptic drive can be estimated from the ratio of the anestheticinduced changes in overall neurotransmission to the changes in the postsynaptic receptor response [94]. The overall excitatory and overall inhibitory drives to respiratory premotor neurons were determined by complete blockade of GABAAergic
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Figure 15.17 (Top) Effect of GABAA receptor activation with muscimol and GABAA receptor block with bicuculline on expiratory neuronal firing pattern. The traces show short record segments of the neuronal response together with the simultaneously recorded phrenic neurogram (PNG). (Bottom) GABAA receptor mediated changes in neuronal activity follow a multiplicative mechanism. During control conditions, the overall excitatory drive to the neuron (Fe) is attenuated by the prevailing GABAergic inhibition ( ) to Fcon (bold). Application of muscimol reduces Fcon to Fmus. The neuronal response to GABAA receptor stimulation () can be expressed as the net decrease from Fcon, normalized to overall excitation. Since the net decrease is calculated from the linear dose-response curve to muscimol, we now use the slope of the dose-response curve, normalized to Fe, for assessment of the anesthetic effect (adapted from Ref. 94).
inhibition with the competitive antagonist bicuculline [93]. The neuronal discharge frequency at complete block of inhibition is equivalent to the overall excitatory drive to the neuron (Fe) (Figure 15.19). The difference between neuronal control frequency (Fcon) and overall excitation (complete GABAergic block) indicates the degree to which Fe is attenuated by the tonic GABAergic inhibition. Overall inhibition ( ) is expressed as the ratio of the physiologic attenuation to overall excitation ( ¼ (FeFcon)/Fe) (Figure 15.17). The postsynaptic receptor response has been determined by picoejection of the respective receptor agonist onto a neuron [30,81,94,95]. Comparison of dose-responses at 0 and 1 MAC anesthesia allowed conclusions about the anesthetic effect on the postsynaptic receptor response. An example, the GABAA receptor antagonist muscimol, is shown in Figure 15.20. Since very small amounts of these agonists produce potent,
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Figure 15.18 Model illustrates the hypothetical functional location of postsynaptic receptors that determine inspiratory neuronal firing frequency (Fn). Tonic and phasic glutamatergic inputs to the dendritic tree elicit a dendritic current (Iden). On its path toward the soma, Iden is attenuated by a tonic GABAAergic input, which leads to a shunting of part of the current ( Iden, where 1 and represents overall tonic inhibition). The remaining current reaches the soma (Isoma ¼ (1 ) Iden) and its magnitude determines neuronal discharge frequency (Fn). There is evidence for the existence of additional glutamate receptors on the soma and proximal dendrites. A possible function of these receptors is the mediation of behavioral inputs like coughing, sneezing or speech, i.e., activities that also require respiratory musculature and can alter spontaneous breathing (adapted from Ref. 95).
rapid changes in neuron discharge frequency, it is likely that these picoejections mainly affect AMPA, NMDA, and GABAA receptors close to the soma [321] with the assumption that those receptors are representative of the receptors located more distally, for example on the peripheral dendrites that are synaptically activated (Figure 15.18). In eight studies on respiratory premotor neurons in the decerebrate in vivo preparation, we sought to answer three main questions: (1) Is the anesthetic depression of premotor neurons predominantly due to an increase in GABAergic inhibition or rather to a depression of glutamatergic excitation? (2) Is there a difference in the anesthetic effect on inspiratory and expiratory premotor neurons? (3) Are the effects of halothane different from those of sevoflurane? Study results indicate that 1 MAC halothane and 1 MAC sevoflurane similarly depress the discharge frequency of respiratory premotor neurons (Figure 15.21A) by depressing overall excitatory drive (Figure 15.21B) and enhancing the GABAA receptor response (Figure 15.21C). The overall inhibition was increased much less than the postsynaptic GABAA receptor response (Figure 15.21C), i.e., the enhanced GABAA receptor response
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Figure 15.19 Dose-response of an inspiratory neuron to the GABAA antagonist bicuculline at 0 and 1 MAC halothane. The picoejection response curves are shown as the rate-meter recordings of the neuronal discharge frequency Fn (in hertz). Bicuculline is ejected in increasing dose-rates (see marker) until an increase in doserate does not lead to a further increase in neuronal discharge activity, i.e., the response is saturated and all GABAA receptors are blocked. At this point, Fn represents the overall excitatory drive to the neuron (Fe). Note the increase in breathing frequency at 1 MAC halothane. (Fcon: neuronal control discharge frequency) (adapted from Ref. 95).
appears to be at least partly offset by an anesthetic depression of presynaptic inhibitory drive. In a previous study in neuraxis-intact animals, we found that an increase in halothane from 1 to 2 MAC actually decreased the overall inhibitory drive by 34% [93]. Similarly, in hippocampal slice CA1 neurons, the enhancement of postsynaptic GABAA-receptor function by volatile anesthetics peaked between 1 and 2 MAC [202]. We suggest that in our in vivo preparation, presynaptic inhibitory drive to the neuron continuously decreased with increasing depth of anesthesia while GABAA receptor function was enhanced with a maximum between 1 and 2 MAC. The resultant overall inhibition was thus increased at low anesthetic levels and decreased at 2 MAC. The most prominent difference between inspiratory and expiratory neurons is that the overall excitatory and inhibitory drives are greater to inspiratory neurons [86,88,95,96] than to expiratory neurons [30,81,94]. In inspiratory neurons, overall inhibition is increased by sevoflurane [88,95] but unchanged by halothane [86,95]. Also, sevoflurane enhances the GABAA-receptor response in inspiratory neurons [88] more than in
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Figure 15.20 (A) Dose-response of an expiratory neuron to increasing dose-rates of the GABAA-agonist muscimol (see marker) at 0 and 1 MAC halothane. The picoejection response curves are shown as the rate-meter recordings of the neuronal discharge frequency Fn (in hertz). Note the much larger depression of neuron activity by muscimol (dotted line ¼ Fcon) at similar maximal dose-rates at 1 MAC halothane. Also note the increase in respiratory rate during anesthesia. (B) Method used to analyze the effect of halothane on postsynaptic GABAA receptor function. The graph shows the response to picoejection of muscimol onto the neuron in (A). Linear regression analysis was performed where the y-intercept was constrained to pass through control frequency (Fcon) at the zero dose-rate. The anesthetic effect in terms of changes in neuronal frequency can be illustrated by interpolation of the net decrease at an identical dose-rate, e.g., at muscimol 0.081 pmol/min, from the doseresponse curve at 0 MAC (solid line) and at 1 MAC (dashed line). Halothane caused an enhancement of the muscimol-induced net decrease from 20 to 49 Hz (adapted from Ref. 94).
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Figure 15.21 Pooled summary data for effects of 2 volatile anesthetics on respiratory premotor neurons. Mean changes SE caused by 1 MAC anesthesia to (A) control frequency (Fcon); (B) overall excitation (Fe); postsynaptic AMPA receptor response (AMPA); postsynaptic NMDA receptor response (NMDA); (C) overall inhibition ( ) and postsynaptic GABAA receptor response (). The effects on inspiratory (I) and expiratory (E) neurons are presented separately. (x: anesthetic effect different from no change with p 5 0.05; *: magnitude of the anesthetic effect different between inspiratory and expiratory neurons with p 5 0.05.)
expiratory neurons [94] (Figure 15.21C). We have no conclusive explanation for this difference, but several possibilities exist. For example, GABAAreceptor density may vary between neuron types. It is also possible that the GABAA receptors on inspiratory neurons consist of a different receptor subtype with greater affinity for GABA. Such a subtype may also have a differential affinity for halothane versus sevoflurane so that receptor
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function is more enhanced with sevoflurane [88]. Alternatively, the conformational changes that result in enhanced receptor function are more pronounced with sevoflurane than with halothane. Sevoflurane and halothane also have different effects on the glutamate receptor responses in inspiratory neurons. The AMPA and NMDA receptor responses are significantly depressed by halothane [95] but not by sevoflurane [96] (Figure 15.21B). This difference may again be explained by the presence of receptor subtypes with differential sensitivity for anesthetics on inspiratory versus expiratory neurons. Alternatively, the depression of the glutamate receptor responses by 1 MAC halothane may indicate that all volatile anesthetics depress glutamate receptor function [180,183] but that the magnitude of depression by 1 MAC of sevoflurane is too small (520%) to be reliably discovered by our method [95]. A 30–40% depression of NMDA and AMPA mediated excitatory currents by enflurane has been convincingly shown for spinal motoneurons [180]. However, contrary results were obtained in a hippocampal slice preparation, where halothane blocked excitatory neurotransmission only at a presynaptic site and did not influence AMPA and NMDA receptors [322].
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In summary, in the in vivo canine decerebrate preparation, 1 MAC halothane and sevoflurane depress neurotransmission to respiratory premotor neurons by a depression of overall excitatory drive and an enhancement of overall inhibition (Figure 15.22). The enhancement of inhibition is due to a prominent increase in postsynaptic GABAA-receptor response, which appears to be partly offset by a decrease in presynaptic net release of GABA. B. Effects of Anesthetics on the Membrane Potential of Respiratory Neurons In vivo
Brainstem respiratory neurons receive both excitatory and inhibitory synaptic inputs that shape the membrane potential trajectory and contribute to the generation of action potentials. When such inputs cause the opening of ligand-gated ion channels, conductivity of the channels increases, which can be measured as a decrease in the membrane input resistance. If the input is excitatory, the membrane potential will become less negative; if inhibitory, the membrane will hyperpolarize. These distinctions can be made with intracellular recordings that allow measurement of the membrane potential and the membrane input resistance. Haji, Takeda and colleagues have performed several difficult studies using intracellular recordings to investigate the in vivo effects of anesthetics and sedatives on bulbar respiratory neurons in a decerebrate cat model. Their setup allows unique insights; however, the technical difficulty of stable recordings limits the investigation to small doses of systemically applied anesthetics or microiontophoretic application of the agents [92,97,98,323]. With either method, the effect site concentrations cannot be reliably estimated. Selective Enhancement of GABAAergic Inhibition and Block of Glutamatergic Excitation
Sedatives, such as the benzodiazepines, are routinely used in combination with other anesthetics and analgesics. The mechanism of action of the benzodiazepines is well established as GABAA-receptor modulators that enhance the effects of GABA. Their specific effects on the membrane potential of bulbar respiratory neurons have been investigated with intracellular recording techniques and microiontophoresis. Iontophoretically-applied flurazepam augmented the IPSPs during the inactive phase of the respiratory cycle (phasic inhibition) of all inspiratory and expiratory neurons [98]. Simultaneously, the membrane input resistance was reduced by 40–60%, which is to be expected since benzodiazepines increase chloride conductance via GABAA-receptor potentiation. Microiontophoretic application of the benzodiazepine consistently hyperpolarized the membrane (3–12 mV), especially during the silent phase of the neuron, but had no effect on EPSPs and did not change the shape and firing threshold of action potentials.
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Figure 15.22 Summary scheme of the effect of 1 MAC halothane (A and C) and sevoflurane (B and C) on inspiratory (A and B) and expiratory (C) premotor neurons in the caudal VRG. The anesthetic effects on excitatory and inhibitory neurotransmission are divided into their presynaptic, postsynaptic and overall components. The combined effects lead to a decrease in control discharge frequency (Fcon). (": increase; ø: no effect; #: decrease) (adapted from Ref. 94, courtesy of Lippincott, Williams & Wilkins, Baltimore, Maryland).
This suggests that any reduction in neuronal discharge frequency is solely due to the relative membrane hyperpolarization. Furthermore, the effects of flurazepam were completely eliminated with microiontophoretic application of the GABAA antagonist bicuculline, suggesting that they were indeed
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Figure 15.22
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GABAA-receptor mediated. Intravenous diazepam exerted qualitatively similar effects on respiratory neurons [97]. Intracellular recordings combined with microiontophoresis that investigate NMDA receptor function in medullary respiratory neurons further verify the validity and usefulness of this technique. That is, the responses are consistent with those expected according to the mechanism of action of the applied receptor agonist or antagonist. For example, antagonism of NMDA receptors with dizocilpine (MK-801) reduces the depolarization of the membrane potential during the active phase together with an increase in input resistance [323] (Figure 15.23). Before
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Figure 15.23 Membrane potential (MP) of an inspiratory neuron and the simultaneously recorded phrenic neurogram (PNG) before and 30 min after systemic administration of the NMDA receptor antagonist dizocilpine. Note the prolongation of inspiratory time (apneusis) and the depression of the neuronal membrane potential with inhibition of action potential discharge (adapted from Ref. 323).
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Effects of Halothane and Thiopental
During intracellular recordings of bulbar respiratory neurons, small doses of halothane (2% for 90 sec by inhalation) or thiopental (2.5 mg/kg i.v.) were systemically given to decerebrate cats and their effects on neuronal membrane fluctuations were analyzed. The most prominent and consistent finding of this study [92] was an increase in membrane resistance that was independent of the type of anesthetic or neuron. This points to a decrease in the level of synaptic input and/or a reduction of postsynaptic receptor function. An anesthetic depression of presynaptic excitatory as well as inhibitory drive has also been shown in reduced preparations [118,324] and postulated for in vivo preparations [30,81,86,88]. This leads to preparations a decrease in excitatory and inhibitory channel opening and thus to an increase in membrane resistance. Any reduction in membrane resistance from the potentiating effect of halothane and thiopental on postsynaptic neuronal GABAA receptors [97] may be offset by such decreases in the level of synaptic input [86,94]. This interpretation is supported by the observed decrease in EPSPs and IPSPs in Takeda’s preparation [92]. Also, the membrane potential fluctuations were reduced. Approximately half the neurons were hyperpolarized while the rest were depolarized. This neuronal response was consistent for both anesthetic agents. Since neurons receive differential synaptic inputs depending on their location in the respiratory network, it is likely that in some neurons the inhibitory drive was more reduced leading to a depolarization of the neuron, while in others the excitatory drive was more reduced resulting in a hyperpolarization of the membrane potential. Membrane hyperpolarization was always accompanied by a decrease in neuronal discharge frequency or the complete cessation of firing as is expected when neuronal excitability is reduced. On the other hand, membrane depolarization of a neuron is expected to increase discharge frequency. This was observed in only a few neurons, but the majority showed a decrease in discharge frequency or even a cessation of firing. The authors suggest that a pronounced depolarization may have uncoupled the steps of action potential generation, i.e., the initial segment spike from the somato-dendritic spike, and thus failed to elicit action potentials [92]. It does not appear that the relative depolarizations are due to direct current drift, since they were consistently observed both for repeated anesthetic applications and for halothane and thiopental within the same neuron. Due to the systemic route of delivery and short duration of application, it might be possible that pharmacokinetics produce transient network conditions that resulted in the observed depolarization coupled with depressed discharge rates. Resolution of this question requires studies with steady-state doses of anesthesia.
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In summary, these difficult studies provide additional evidence that the respiratory depression by halothane and thiopental in vivo is due to several mechanisms. Both anesthetics depressed excitatory and inhibitory drive to bulbar respiratory neurons. These anesthetic effects are clearly different from agents that selectively modulate only one type of receptor. C. Anesthetic Depression of Hypoglossal Motoneurons: Role of Anesthetic-induced KQ Channels
Respiratory-related activity is also found in HMs that innervate the tongue muscles. Inspiratory hypoglossal activity becomes more prominent during hypercapnia and other states of increased respiratory drive. The tongue plays a crucial role in upper airway patency, and decreased activity of the HM has been related to obstructive sleep apnea [325]. Berger, Bayliss and their collaborators have extensively researched the physiology of HMs with in vitro electrophysiological recordings in brainstem slice preparations [162]. The neuronal firing pattern and the composition of receptors on the HM neuronal cell membrane change greatly during maturation from the neonatal to the adult rat [326,327]. Here we will focus on the adult animal. Similar to other respiratory motoneurons, the primary excitatory drive to HMs is glutamatergic and mediated mostly by non-NMDA receptors, while NMDA receptor activation may regulate the excitability of the cells [328]. Both glutamate receptor types can be co-localized in a single synapse [329]. HMs are also subject to inhibitory inputs mediated by GABA and glycine, which are co-released from presynaptic vesicles [330]. Unlike other respiratory motoneurons, inspiratory HMs are not periodically inhibited during the expiratory phase but only during the later part of the inspiratory cycle. This inhibition limits the inspiratory phase depolarization, which helps turn off the neuron [331]. Qian and Sejnowski observed that the inhibitory input leads to an increase in membrane conductance without a significant change in membrane potential [332]. They conclude that this type of inhibition, which is localized proximal, i.e., closer to the soma, than the excitatory input, allows shunting of the excitatory input and very effectively reduces excitation. Zuperku et al. have suggested a similar mechanism for respiratory premotor neurons; they observed a gain modulatory effect of GABAergic inhibition on the respiratory firing pattern of premotor neurons [296] (for details see section V.A). Modulation of Hypoglossal Motoneuron Excitability
The excitability of HMs is modulated by the neurotransmitters serotonin (5-HT), norepinephrine (NE), thyrotropin releasing hormone (TRH), substance P (SP), and glutamate, when they bind to their specific receptors, i.e., the 5-HT2, a1, TRH-R1, NK1, and metabotropic glutamate receptors, respectively [333] (Figure 15.24). All these receptors are
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Figure 15.24 Diagram of ligand-mediated modulation of hypoglossal motoneuronal excitability and of the effects of acidosis and volatile anesthetics on neuronal background Kþ channels. Binding of a ligand to the Gaq-protein coupled receptor elicits a response via second messengers that leads to an increased opening of bariumresistant Naþ channels and a depression of barium-sensitive leak Kþ channels. Both effects result in a depolarization of the cell. The leak Kþ channel can also be blocked by a decrease in extracellular pH. In contrast, volatile anesthetics open the channel and lead to a hyperpolarization of the cell and thus decrease neuronal excitability (adapted from Ref. 333, courtesy of Elsevier, Oxford, UK).
G-protein coupled and use only partially identified second-messenger pathways to increase the excitability of the cell by depolarization [327]. The depolarization appears to result from the activation of barium-resistant Naþ channels and from a block of barium-sensitive resting (leak) Kþ channels. Block of this Kþ channel leads to an increase in neuronal input resistance (RN) and results in a depolarization of the membrane so that less excitatory current is required to reach firing threshold. At the same time, an increased RN also amplifies the membrane voltage response to inhibitory synaptic currents. Thus, the binding of these neurotransmitters does not affect the inspiratory and expiratory phase duration, but amplifies the postsynaptic response to excitatory and inhibitory inputs [327]. The channel most likely responsible for the neuromodulator-mediated depolarization of HMs has been identified as the TASK-1 (Tandem pore Acid Sensitive Kþ) channel, a leak Kþ channel with four transmembrane
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regions and two pore domains; the channel is acid-sensitive and modulated in the physiologic pH range (pK 7.4) [333]. TASK-1 channels have been found in great numbers on brainstem and spinal cord motoneurons, including HMs [162]. In a rat brainstem slice preparation with HMs under current clamp conditions, acidification of the bath solution (pH 6.5) leads to a depolarization of the membrane until it reaches the threshold for repetitive discharge [333]. On the other hand, alkalization of the bath solution (pH 8.4) hyperpolarizes the membrane [162]. Neurotransmitters, e.g., 5-HT, completely block the leak current even when it is enhanced by high pH [162] (Figure 15.25). These effects are relevant in the physiologic pH range, i.e., a shift in extracellular pH from 7.4 to 7.1 is associated with a 10% reduction in input conductance [333], and suggests a role for the TASK-1 channel in mediating chemoreception [333]. The presence of TASK-1 channels has also been shown on putative chemoreceptor neurons in the locus coeruleus [164] and raphe´ [334]. An extracellular decrease in pH may thus facilitate depolarization of chemoreceptive cells by inactivation of TASK channels. Interestingly, the TASK-1 channel is activated by inhalational anesthetics (see below). This activation may counteract the excitatory effect of low pH on the neurons and may be one mechanism by which inhalational anesthetics reduce the central chemodrive to the respiratory system. The relevance of a chemoreceptive capacity for respiratory motoneurons has to be assessed in an in vivo model, since in neurons that are subject to inhibitory inputs during their active phase, e.g., phrenic and external intercostal motoneurons [335], an increase in RN would not necessarily result in a depolarization of the cells. Activation of TASK-1 Channels by Inhalational Agents
Using a rat brainstem slice preparation, Sirois et al. found that bath application of halothane and sevoflurane in clinically relevant concentrations causes an outward current in voltage-clamped HMs [164]. The current is dose-dependent and can be blocked by low extracellular pH [164] (Figure 15.24). A similar effect on TASK channels has been shown for isoflurane [14]. Halothane also inhibits a hyperpolarization-induced Naþ current and a TTX-insensitive Naþ current in HMs, but activation of TASK channels produces a greater degree of hyperpolarization [100]. TASK channels have also been found on spinal motoneurons [162], though not explicitly on the phrenic motoneurons. If TASK channels are present on phrenic motoneurons, this may explain the greater degree of anestheticinduced depression of peak phrenic nerve activity relative to that seen for respiratory premotor neurons, on which TASK channels do not appear to be present in the in vivo dog model [30].
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Figure 15.25 The motoneuronal pH-sensitive current is modulated by multiple neurotransmitters. (A) Left, time series showing the effect of serotonin (5-HT) on holding current of hypoglossal motoneuron (HM) in control solution (pH 7.3), and in solutions of pH 6.5 and pH 8.4. The response to 5-HT was first reduced and then enhanced by, respectively, inhibition and activation of the pH-sensitive Kþ conductance. Right, averaged data for HMs treated with 5-HT in control solution and subsequently in pH 6.5 and/or pH 8.4. In all cases the response to 5-HT was smaller following acidification and larger following alkalization. (*: p 5 0.05 by paired t-test.) (B) HMs were exposed to a low pH solution (pH 6.5) prior to and following bath application of neurotransmitters, as follows: 5-HT, norepinephrine (NE), substance P (SP), and thyrotropin-releasing hormone (TRH). Note that the pH-sensitive current at 60 mV was completely blocked by neurotransmitter in each of these cells (adapted from Ref. 333, courtesy of Elsevier, Oxford, UK).
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Summary and Outlook
The depressant effect of general anesthetics on the central respiratory network appears to be mediated by a few major mechanisms. The primary neuronal targets include the ligand-gated channels involved in fast synaptic neurotransmission, anesthesia-activated background Kþ channels and, possibly, voltage-gated channels involved in presynaptic neurotransmitter release. A small number of in vivo studies in respiratory neurons have been very useful in assessing the relative importance of these anesthetic mechanisms on synaptic transmission in the functioning respiratory network. These studies show that the anesthetic depression of the respiratory network is not due to a singular mechanism, but affects both excitatory and inhibitory neurotransmission. In addition, in vivo studies on respiratory premotor neurons suggest that volatile anesthetics affect neurotransmission at the presynaptic and, at least for the GABAA receptor, at the postsynaptic level. These findings have to be considered in the development of anesthetic drugs with less respiratory side effects. Other components of the respiratory network whose physiology has been detailed in this chapter have not yet been investigated for the effects of anesthesia. One important future focus is on the chemoreceptive neurons that mediate respiratory drive and on the neurons involved in rhythm and pattern generation that are located in the preBo¨tzinger complex. Another important area is phrenic, and in particular cranial, respiratory motoneurons involved in airway patency. On the latter neurons, anesthesiasensitive potassium channels have been demonstrated in vitro. The importance of this effect for neuronal activity in the in vivo network, however, remains to be established.
Acknowledgment Work in this laboratory was supported by grant No. GM59234-01, ‘Volatile Anesthetics and Respiratory Neurotransmission’ from the National Institutes of Health, Bethesda, Maryland (to Dr. Stuth), and VA Medical Research Funds, Washington, DC (to Dr. Zuperku).
References 1.
Lumb, A.B., Anesthesia, Anonymous, in Nunn’s Applied Respiratory Physiology, 5th edn., Oxford, Butterworth-Heinemann, pp. 420–452, 2000. 2. Pavlin, E.G. and Hornbein, T.F., The respiratory system, in Handbook of Physiology, Geiger, S.R., ed., Baltimore, The Williams & Wilkins Company, pp. 793–813, 1986.
Central Effects of General Anesthesia 3.
4. 5.
6. 7.
8.
9.
10.
11.
12.
13.
14.
15. 16.
17.
18.
631
Farber, N.E., Pagel, P.S. and Warltier, D.C., Pulmonary pharmacology, in Miller’s Anesthesia, Miller, R.D., ed., 5th ed., New York, Churchill Livingstone, pp. 125–161, 2000. Eger, E., A brief history of the origin of minimum alveolar concentration (MAC), Anesthesiology 96, 238–239, 2002. Eger, E., II, Saidman, L.J. and Brandstater, B., Minimum alveolar anesthetic concentration: A standard of anesthetic potency, Anesthesiology 26, 756–763, 1965. Rampil, I.J. and King, B.S., Volatile anesthetics depress spinal motor neurons, Anesthesiology 85, 129–134, 1996. Antognini, J.F., Carstens, E., Tabo, E. and Buzin, V., Effect of differential delivery of isoflurane to head and torso on lumbar dorsal horn activity, Anesthesiology 88, 1055–1061, 1998. Knill, R.L. and Clement, J.L., Site of selective action of halothane on theperipheral chemoreflex pathway in humans, Anesthesiology 61, 121–126, 1984. Dwyer, R., Benett, H.L., Eger, E., 2nd and Heilbron, D., Effects of isoflurane and nitrous oxide in subanesthetic concentrations on memory and responsiveness in volunteers, Anesthesiology 77, 888–898, 1992. Nandi, P.R., Charlesworth, C.H., Taylor, S.J., Nunn, J.F. and Dore, C.J., Effect of general anaesthesia on the pharynx, Br. J. Anaesth. 66, 157– 162, 1991. Hwang, J.-C., St. John, W.M. and Bartlett, D., Jr., Respiratory-related hypoglossal nerve activity: influence of anesthetics, J. Appl. Physiol. 55, 785–792, 1983. Nishino, T., Kohchi, T., Yonezawa, T. and Honda, Y., Responses of recurrent laryngeal, hypoglossal, and phrenic nerves to increasing depths of anesthesia with halothane or enflurane in vagotomized cats, Anesthesiology 63, 404–409, 1985. Ochiai, R., Guthrie, R.D. and Motoyama, E.K., Effects of varying concentrations of halothane on the activity of the genioglossus, intercostals, and the diaphragm in cats: An electromyographic study, Anesthesiology 70, 812–816, 1989. Patel, A.J., Honore, E., Lesage, F., Fink, M., Romey, G. and Lazdunski, M., Inhalational anesthetics activate two-pore-domain background Kþ channels, Nat. Neurosci. 2, 422–426, 1999. Nattie, E., CO2, brainstem chemoreceptors and breathing, Prog. Neurobiol. 59, 299–331, 1999. Munson, E.S., Larson, C.P., Bobad, A.A., Regan, M.J., Buechel, D.R. and Eger, E., II, The effects of halothane, flurane and cyclopropane on ventilation: A comparative study in man, Anesthesiology 27, 716–728, 1966. Lam, A.M., Clement, J.L., Chung, D.C. and Knill, R.L., Respiratory effects of nitrous during enflurane anesthesia in humans, Anesthesiology 56, 298–303, 1982. Calverley, R.K., Smith, N.T., Jones, C.W., Prys-Roberts, C. and Eger, E.I., 2nd, Ventilatory and cardiovascular effects of enflurane anesthesia during spontaneous ventilation in man, Anesth. Analg. 57, 610–618, 1978.
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31. 32.
33.
Stuth et al. Lockhart, S.H., Rampil, I.J., Yasuda, N., Eger, E.I., 2nd, Weiskopf, R.B., Depression of ventilation by desflurane in humans, Anesthesiology 74, 484–488, 1991. Eger, E., II, Isoflurane: A review, Anesthesiology 55, 559–576, 1981. Fourcade, H.E., Stevens, W.C., Larson, C.P., Jr., Cromwell, T.H., Bahlman, S.H., Hickey, R.F., Halsey, M.J., Eger, E.I., 2nd, The ventilatory effects of Forane, a new inhaled anesthetic, Anesthesiology 35, 26–31, 1971. Doi, M. and Ikeda, K., Respiratory effects of sevoflurane, Anesth. Analg. 66, 241–244, 1987. Green, W.B., Jr., The ventilatory effects of sevoflurane, Anesth. Analg. 81, S23–S26, 1995. Dahan, A., Olievier, I.C.W., Berkenbosch, A. and DeGoede, J., Modeling the dynamic ventilatory response to carbon dioxide in healthy human subjects during normoxia, in Respiratory Control, Swanson, G.D., Grodins, F.S. and Hughson, R.L., eds., New York, Plenum Publishing Corporation, pp. 265–273, 1989. Berkenbosch, A., Dahan, A., DeGoede, J. and Olievier, I.C.W., The ventilatory response to CO2 of the peripheral and central chemoreflex loop before and after sustained hypoxia in man, J. Physiol. (Lond.) 456, 71–83, 1992. Knill, R.L. and Gelb, A.W., Ventilatory responses to hypoxia and hypercapnia during halothane sedation and anesthesia in man, Anesthesiology 49, 244–251, 1978. Knill, R.L., Manninen, P. and Clement, J.L., Ventilation and chemoreflexes during enflurane sedation and anesthesia in man, Can. Anaesth. Soc. J. 26, 353– 360, 1979. Stuth, E.A.E., Tonkovic-Capin, M., Kampine, J.P. and Zuperku, E.J., Dose-dependent effects of isoflurane on the CO2 responses of expiratory medullary neurons and the phrenic nerve activities in dogs, Anesthesiology 76, 763–774, 1992. Stuth, E.A.E., Tonkovic-Capin, M., Kampine, J.P., Bajic, J. and Zuperku, E.J., Dose-dependent effects of halothane on the carbon dioxide responses of expiratory and inspiratory bulbospinal neurons and the phrenic nerve activities in dogs, Anesthesiology 81, 1470–1483, 1994. Stucke, A.G., Stuth, E.A.E., Tonkovic-Capin, V., Tonkovic-Capin, M., Hopp, F.A., Kampine, J.P. and Zuperku, E.J., Effects of sevoflurane on excitatory neurotransmission to medullary expiratory neurons and on phrenic nerve activity in a decerebrate dog model, Anesthesiology 95, 485– 491, 2001. Eldridge, F.L., Relationship between respiratory nerve and muscle activity and muscle force output, J. Appl. Physiol. 39, 567–574, 1975. Fierobe, L., Cantineau, J.P., Pandele, G. and Desmonts, J.M., Measurement of respiratory effects of propofol by indirect spirometry, Ann. Fr. Anesth. Reanim. 10, 10–15, 1991. Tusiewicz, K., Bryan, A.C. and Froese, A.B., Contributions of changing rib cage–diaphragm interactions to the ventilatory depression of halothane anesthesia, Anesthesiology 47, 327–337, 1977.
Central Effects of General Anesthesia 34.
35.
36.
37.
38. 39. 40. 41. 42. 43.
44.
45. 46.
47.
48.
49.
633
Allsop, P., Taylor, M.B., Grounds, R.M. and Morgan, M., Ventilatory effects of a propofol infusion using a method to rapidly achieve steady-state equilibrium, Eur. J. Anaesthesiol. 5, 293–303, 1988 (Comment in 6, 397–398, 1989). Nieuwenhuijs, D., Sarton, E., Teppema, L.J., Kruyt, E., Olievier, I., vanKleef, J. and Dahan, A., Respiratory sites of action of propofol: Absence of depression of peripheral chemoreflex loop by low-dose propofol, Anesthesiology 95, 889–895, 2001. Yamakage, M., Kamada, Y., Toriyabe, M., Honma, Y. and Namiki, A., Changes in respiratory pattern and arterial blood gases during sedation with propofol or midazolam in spinal anesthesia, J. Clin. Anesth. 11, 375–379, 1999. Reber, A., Wetzel, S.G., Schnabel, K., Bongartz, G. and Frei, F.J., Effect of combined mouth closure and chin lift on upper airway dimensions during routine magnetic resonance imaging in pediatric patients sedated with propofol, Anesthesiology 90, 1617–1623, 1999. Kohrs, R. and Durieux, M.E., Ketamine: Teaching an old drug new tricks, Anesth. Analg. 87, 1186–1193, 1998. Liburn, J.K., Dundee, J.W. and Moore, J., Ketamine infusions, Anaesthesia 33, 315–321, 1978. Idvall, J., Ahlgren, I., Aronsen, K.F. and Stenberg, P., Ketamine infusion: Pharmacokinetics and clinical effects, Br. J. Anaesth. 51, 1167–1173, 1979. Hamza, J., Ecoffey, C. and Gross, J.B., Ventilatory response to CO2 following intravenous ketamine in children, Anesthesiology 70, 422–425, 1989. Bourke, D., Malit, L.A. and Smith, T.C., Respiratory interactions of ketamine and morphine, Anesthesiology 66, 153–156, 1987. Sarton, E., Teppema, L.J., Olievier, C., Nieuwenhuijs, D., Matthes, H.W.D., Kieffer, B.L. and Dahan, A., The involvement of the m-opioid receptor in ketamine-induced respiratory depression and antinociception, Anesth. Analg. 93, 1495–1500, 2001. Shulman, D., Bar-Yishay, E. and Godfrey, S., Drive timing components of respiration in young children following induction of anaesthesia with halothane or ketamine, Can. J. Anaesth. 35, 368–374, 1988. Jaspar, N., Mazzarelli, M., Tessier, C. and Milic-Emili, J., Effect of ketamine on control of breathing in cats, J. Appl. Physiol. 55, 851–859, 1983. Pierrefiche, O., Foutz, A.S. and Denavit-Saubie´, M., Pneumotaxic mechanisms in the non-human primate: Effect of the N-methyl-D-aspartate (NMDA) antagonist ketamine, Neurosci. Lett. 119, 90–93, 1990. Pierrefiche, O., Foutz, A.S., Champagnat, J. and Denavit-Saubie´, M., NMDA and non-NMDA receptors may play distinct roles in timing mechanisms and transmission in the feline respiratory network, J. Physiol. (Lond.) 474, 509–523, 1994. Foutz, A.S., Pierrefiche, O. and Denavit-Saubie´, M., Combined blockade of NMDA and non-NMDA receptors produces respiratory arrest in the adult cat, Neuroreport 5, 481–484, 1994. Pokorski, M., Sieradzan, B. and Karczewski, W., Apneustic respiration of ketamine is not antagonized in the cat, Jap. J. Physiol. 37, 735–740, 1987.
634 50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62. 63.
64.
Stuth et al. Foutz, A.S., Champagnat, J. and Denavit-Saubie´, M., N-methyl-D-aspartate (NMDA) receptors control respiratory off-switch in cat, Neurosci. Lett. 87, 221–226, 1988. Shulman, D., Bar-Yishay, E., Beardsmore, C. and Godfrey, S., Determinants of end expiratory volume in young children during ketamine or halothane anesthesia, Anesthesiology 66, 636–640, 1987. Shulman, D.L., Bar-Yishay, E. and Godfrey, S., Respiratory mechanics and intrinsic PEEP during ketamine and halothane anesthesia in young children, Anesth. Analg. 67, 656–662, 1988. Shulman, D., Beardsmore, C.S., Aronson, H.B. and Godfrey, S., The effect of ketamine on the functional residual capacity in young children, Anesthesiology 62, 551–556, 1985. Mankikian, B., Cantineau, J.P., Sartene, R., Clergue, F. and Viars, P., Ventilatory pattern and chest wall mechanics during ketamine anesthesia in humans, Anesthesiology 65, 492–499, 1986. Warner, D.O., Warner, M.A. and Ritman, E.L., Human chest wall function while awake and during halothane anesthesia. I. Quiet breathing, Anesthesiology 82, 6–19, 1995. Warner, D.O. and Warner, M.A., Human chest wall function while awake and during halothane anesthesia. II. Carbon dioxide rebreathing, Anesthesiology 82, 20–31, 1995. Hirshman, C.A., McCullough, R.E., Cohen, P.J. and Weil, J.V., Hypoxic ventilatory drive in dogs during thiopental, ketamine, or pentobarbital anesthesia, Anesthesiology 43, 628–634, 1975. Hirshman, C.A., McCullough, R.E., Cohen, P.J. and Weil, J.V., Depression of hypoxic ventilatory response by halothane, enflurane and isoflurane in dogs, Br. J. Anaesth. 43, 957–963, 1977. Gautier, H., Offenstadt, G., Kaczmarek, R., Bonora, M., Pinta, P. and Hericoard, P., Pattern of respiration in patients recovering from barbiturate overdose, Br. J. Anaesth. 54, 1041–1045, 1982. Launois, S., Similowski, T., Fleury, B., Aubier, M., Murciano, D., Housset, B., Pariente, R. and Derenne, J.P., The transition between apnoea and spontaneous ventilation in patients with coma due to voluntary intoxication with barbiturates and carbamates, Eur. Respir. J. 3, 573–578, 1990. Siafakas, N.M., Bonora, M., Duron, B., Gautier, H. and Milic-Emili, J., Dose effect of pentobarbital sodium on control of breathing in cats, J. Appl. Physiol. 55, 1582–1592, 1983. Gautier, H., Pattern of breathing during hypoxia or hypercapnia of the awake or anesthetized cat, Respir. Physiol. 27, 193–206, 1976. Gross, J.B., Zebrowski, M.E., Carel, W.D., Gardner, S. and Smith, T.C., Time course of ventilatory depression after thiopental and midazolam in normal subjects and in patients with chronic obstructive pulmonary disease, Anesthesiology 58, 540–544, 1983. Saetta, M. and Mortola, J.P., Breathing pattern and CO2 response in newborn rats before and during anesthesia, J. Appl. Physiol. 58, 1988–1996, 1985.
Central Effects of General Anesthesia 65.
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
77. 78.
79.
635
Haji, A., Okazaki, M., Yamazaki, H. and Takeda, R., Physiological properties of late inspiratory neurons and their possible involvement in inspiratory off-switching in cats, J. Neurophysiol. 87, 1057–1067, 2002. Gage, P.W. and Robertson, B., Prolongation of inhibitory postsynaptic currents by pentobarbitone, halothane and ketamine in CA1 pyramidal cells in rat hippocampus, Br. J. Pharmacol. 85, 675–681, 1985. Pistis, M., Belelli, D., Peters, J.A. and Lambert, J.J., The interaction of general anaesthetics with recombinant GABAA and glycine receptors expressed in Xenopus laevis oocytes: A comparative study, Br. J. Pharmacol. 122, 1707– 1719, 1997. Wu, J., Harata, N. and Akaike, N., Potentiation by sevoflurane of the g-aminobutyric acid-induced chloride current in acutely dissociated CA1 pyramidal neurones from rat hippocampus, Br. J. Pharmacol. 119, 1013–1021, 1996. Charlesworth, P., Jacobson, I. and Richards, C.D., Pentobarbitone modulation of NMDA receptors in neurones isolated from the rat olfactory brain, Br. J. Pharmacol. 116, 3005–3013, 1995. Dildy-Mayfield, J.E., Eger, E.I., II and Harris, R.A., Anesthetics produce subunit-selective actions on glutamate receptors, J. Pharmacol. Exp. Ther. 276, 1058–1065, 1996. Gautier, M. and Gaudy, F.M., Changes in ventilatory pattern induced by intravenous anesthetic agents in human subjects, J. Appl. Physiol. 45, 171–176, 1978. Marsh, H.M., Rehder, K. and Hyatt, R.E., Respiratory timing and depth of breathing in dogs anesthetized with halothane or enflurane, J. Appl. Physiol. 51, 19–25, 1981. Mazzarelli, M.S., Haberer, F.P., Jaspar, N. and Miserocchi, G., Mechanism of halothane-induced tachypnea in cats, Anesthesiology 51, 522– 527, 1979. Berkenbosch, A., de Goede, J., Olievier, C.N. and Quanjer, P.H., Sites of action of halothane on respiratory pattern and ventilatory response to CO2 in cats, Anesthesiology 57, 389–398, 1982. Paskin, S., Skovsted, P. and Smith, T.C., Failure of the Hering-Breuer reflex to account for tachypnea in anesthetized man, Anesthesiology 29, 550–558, 1968. Grelot, L. and Bianchi, A.L., Differential effects of halothane anesthesia on the pattern of discharge of inspiratory and expiratory neurons in the region of the retrofacial nucleus, Brain Res. 404, 335–338, 1987. Nishino, T. and Honda, Y., Changes in the respiratory pattern induced by halothane in the cat, Br. J. Anaesth. 52, 1191–1197, 1980. Nishino, T., Honda, Y. and Yonezawa, T., Separate effects of halothane and carbon dioxide on respiratory duration in vagotomized cats, Br. J. Anaesth. 55, 647–653, 1983. Mutoh, T., Tsubone, H., Nishimura, R. and Sasaki, N., Effects of volatile anesthetics on vagal C-fiber activities and their reflexes in anesthetized dogs, Respir. Physiol. 112, 253–264, 1998.
636 80. 81.
82. 83. 84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94.
Stuth et al. Gautier, H., Bonora, M. and Zaoui, D., Influence of halothane on control of breathing in intact and decerebrated cats, J. Appl. Physiol. 63, 546–553, 1987. Stuth, E.A.E., Krolo, M., Stucke, A.G., Tonkovic-Capin, M., Tonkovic-Capin, V., Hopp, F.A., Kampine, J.P. and Zuperku, E.J., Effects of halothane on excitatory neurotransmission to medullary expiratory neurons in a decerebrate dog model, Anesthesiology 93, 1474–1481, 2000. Cohen, M.I., Neurogenesis of respiratory rhythm in the mammal, Physiol. Rev. 59, 1105–1173, 1979. Grunstein, M.M., Younes, M. and Milic-Emili, J., Control of tidal volume and respiratory frequency in anesthetized cats, J. Appl. Physiol. 35, 463–476, 1973. Gautier, H., Bonora, M. and Gaudy, J.H., Breuer-Hering inflation reflex and breathing pattern in anesthetized humans and cats, J. Appl. Physiol. 51, 1162–1168, 1981. Doi, K., Kasaba, T. and Kosaka, Y., A comparative study of the depressive effects of halothane and sevoflurane on medullary respiratory neurons in cats, Masui 37, 1466–1477, 1988. Stucke, A.G., Stuth, E.A.E., Tonkovic-Capin, V., Kampine, J.P. and Zuperku, E.J., Effects of halothane on inhibitory neurotransmission to medullary inspiratory neurons in a decerebrate dog model, Anesthesiology A1362, 2002. Smith, J.C., Butera, R.J., Koshiya, N., Del Negro, C., Wilson, C.G. and Johnson, S.M., Respiratory rhythm generation in neonatal and adult mammals: The hybrid pacemaker-network model, Respir. Physiol. 122, 131–147, 2000. Stucke, A.G., Stuth, E.A.E., Krolo, M., Kampine, J.P. and Zuperku, E.J., Sevoflurane enhances overall inhibitory drive and the postsynaptic GABAA receptor response in inspiratory neurons in a decerebrate dog model, FASEB J. 302.3, 2003. Kasaba, T., Kosaka, Y. and Doi, K., A comparative study of the depressive effects of halothane and enflurane from 1 MAC to 3 MAC on medullary respiratory neuron in cats, Masui 36, 1596–1601, 1987. Tabatabai, M., Kitahata, L.M., Yuge, O., Matsumoto, M. and Collins, J.G., Effects of halothane on medullary inspiratory neurons of the cat, Anesthesiology 66, 176–180, 1987. Tabatabai, M., Kitahata, L.M., Kawahara, M. and Collins, J.G., Enflurane depresses activity of the medullary inspiratory neurons in the cat, Proc. Soc. Exp. Biol. Med. 195, 79–83, 1990. Takeda, R., Haji, A., Hukuhara, T., Selective actions of anesthetic agents on membrane potential trajectory in bulbar respiratory neurons of cats, Pflu¨gers Arch. 416, 375–384, 1990. Stuth, E.A.E., Krolo, M., Tonkovic-Capin, M., Hopp, F.A., Kampine, J.P. and Zuperku, E.J., Effects of halothane on synaptic neurotransmission to medullary expiratory neurons in the ventral respiratory group of dogs, Anesthesiology 91, 804–814, 1999. Stucke, A.G., Stuth, E.A.E., Tonkovic-Capin, V., Tonkovic-Capin, M., Hopp, F.A., Kampine, J.P. and Zuperku, E.J., Effects of halothane and sevoflurane on inhibitory neurotransmission to medullary expiratory neurons in a decerebrate dog model, Anesthesiology 96, 955–962, 2002.
Central Effects of General Anesthesia 95.
96.
97.
98.
99.
100.
101. 102. 103.
104.
105. 106. 107. 108. 109. 110.
111.
637
Stucke, A.G., Zuperku, E.J., Tonkovic-Capin, V., Tonkovic-Capin, M., Hopp, F.A., Kampine, J.P. and Stuth, E.A., Halothane depresses glutamatergic neurotransmission to brainstem inspiratory premotor neurons in a decerebrate dog model, Anesthesiology 98, 897–905, 2003. Stucke, A.G., Stuth, E.A.E., Tonkovic-Capin, V., Kampine, J.P. and Zuperku, E.J., Sevoflurane depresses overall excitatory drive but not the postsynaptic glutamate receptor response to medullary inspiratory neurons in a decerebrate dog model, FASEB J. 302.2, 2003. Takeda, R., Haji, A. and Hukuhara, T., Diazepam potentiates postsynaptic inhibition in bulbar respiratory neurons of cats, Respir. Physiol. 77, 173–186, 1989. Takeda, R. and Haji, A., Microiontophoresis of flurazepam on inspiratory and postinspiratory neurons in the ventrolateral medulla of cats: An intracellular study in vivo, Neurosci. Lett. 102, 261–267, 1989. Takeda, R. and Haji, A., Effects of halothane on membrane potential and discharge activity in pairs of bulbar respiratory neurons of decerebrate cats, Neuropharmacology 31, 1049–1058, 1992. Sirois, J.E., Pancrazio, J.J., Lynch, C., III and Bayliss, D.A., Multiple ionic mechanisms mediate inhibition of rat motoneurones by inhalation anaesthetics, J. Physiol. (Lond.) 512, 851–862, 1998. Urban, B.W., Current assessment of targets and theories of anaesthesia, Br. J. Anaesth. 89, 167–183, 2002. Franks, N.P. and Lieb, W.R., Which molecular targets are most relevant to general anaesthesia? Toxicol. Lett. 100–101, 1–8, 1998. Dong, X.P. and Xu, T.L., The action of propofol on gamma-aminobutyric acid-A and glycine receptors in acutely dissociated spinal dorsal horn neurons of the rat, Anesth. Analg. 95, 907–914, 2002. Meir, A., Ginsburg, S., Butkevich, A., Kachalsky, S.G., Kaiserman, I., Ahdut, R., Demirgoren, S. and Rahamimoff, R., Ion channels in presynaptic nerve terminals and control of transmitter release, Physiol. Rev. 79, 1019–1088, 1999. Moreno, D.H., Molecular and functional diversity of voltage-gated calcium channels, Ann. NY Acad. Sci. 868, 102–117, 1999. Catterall, W.A., Interactions of presynaptic Ca2þ channels and snare proteins in neurotransmitter release, Ann. NY Acad. Sci. 868, 144–159, 1999. Catterall, W.A., Structure and function of neuronal Ca2þ channels and their role in neurotransmitter release, Cell Calcium 24, 307–323, 1998. Catterall, W.A., From ionic currents to molecular mechanisms: The structure and function of voltage-gated sodium channels, Neuron 26, 13–25, 2000. Catterall, W.A., Molecular properties of brain sodium channels: An important target for anticonvulsant drugs, Adv. Neurol. 79, 441–456, 1999. Schlame, M. and Hemmings, H.C., Inhibition by volatile anesthetics of endogenous glutamate release from synaptosomes by a presynaptic mechanism, Anesthesiology 82, 1406–1416, 1995. Kendall, T.J. and Minchin, M.C., The effects of anaesthetics on the uptake and release of amino acid neurotransmitters in thalamic slices, Br. J. Pharmacol. 75, 219–227, 1982.
638
Stuth et al.
112. Minchin, M.C., The effect of anaesthetics on the uptake and release of gamma-aminobutyrate and D-aspartate in rat brain slices, Br. J. Pharmacol. 73, 681–689, 1981. 113. Kullmann, D.M., Martin, R.L. and Redman, S.J., Reduction by general anaesthetics of group Ia excitatory postsynaptic potentials and currents in the cat spinal cord, J. Physiol. (Lond.) 412, 277–296, 1989. 114. Weakly, J.N., Effect of barbiturates on ‘quantal’ synaptic transmission in spinal motoneurones, J. Physiol. (Lond.) 204, 63–77, 1969. 115. MacIver, M.B., Mikulec, A.A., Amagasu, S.M. and Monroe, F.A., Volatile anesthetics depress glutamate transmission via presynaptic actions, Anesthesiology 85, 823–834, 1996. 116. Richards, C.D., Anaesthetic modulation of synaptic transmission in the mammalian CNS, Br. J. Anaesth. 89, 79–90, 2002. 117. Westphalen, R.I. and Hemmings, H.C., Selective depression by isoflurane and propofol of glutamate vs. GABA release from isolated cortical nerve terminals, Anesthesiology A-818, 2002. 118. Perouansky, M., Kirson, E.D. and Yaari, Y., Halothane blocks synaptic excitation of inhibitory interneurons, Anesthesiology 85, 1431–1438, 1996. 119. Pierrefiche, O., Haji, A., Bischoff, A. and Richter, D.W., Calcium currents in respiratory neurons of the cat in vivo, Pflu¨gers Arch., 438, 817–826, 1999. 120. Lynch, C., Voltage-gated calcium channels, in Anesthesia: Biologic Foundations, Yaksh, T.L., Lynch, C., Zapol, W.M., Maze, M., Biebuyck, J.F. and Saidman, L.J., eds., Philadelphia, Lippincott-Raven Publishers, pp. 163– 196, 1997. 121. Brevi, S., de Curtis, M. and Magistretti, J., Pharmacological and biophysical characterization of voltage-gated calcium currents in the endopiriform nucleus of the guinea pig, J. Neurophysiol. 85, 2076–2087, 2001. 122. Hofmann, F., Lacinova, L. and Klugbauer, N., Voltage-dependent calcium channels: From structure to function, Rev. Physiol. Biochem. Pharmacol. 139, 33–87, 1999. 123. Catterall, W.A., Structure and regulation of voltage-gated Ca2þ channels, Annu. Rev. Cell Dev. Biol. 16, 521–555, 2000. 124. Heady, T.N., Gomora, J.C., Macdonald, T.L. and Perez-Reyes, E., Molecularpharmacology of T-type Ca2þ channels, Jap. J. Pharmarcol. 85, 339–350, 2001. 125. Suzuki, S. and Rogawaski, M.A., T-type calcium channels mediate the transition between tonic and phasic firing in thalamic neurons, Proc. Natl. Acad. Sci. USA 86, 7228–7732, 1989. 126. White, G., Lovinger, D.M. and Weight, F.F., Transient low-threshold Ca2þ current triggers burst firing through an afterdepolarizing potential in adult mammalian neuron, Proc. Natl. Acad. Sci. USA 86, 6802–6806, 1989. 127. Gomora, J.C., Daud, A.N., Weiergraber, M. and Perez-Reyes, E., Block of cloned human T-type calcium channels by succinimide antiepileptic drugs, Mol. Pharmacol. 60, 1121–1132, 2001. 128. Kavalali, E.T. and Plummer, M.R., Multiple voltage-dependent mechanisms potentiate calcium channel activity in hippocampal neurons, J. Neurosci. 16, 1072–1082, 1996.
Central Effects of General Anesthesia
639
129. Mironov, S.L. and Richter, D.W., Hypoxic modulation of L-type Ca2þ channels in inspiratory brainstem neurones: Intracellular signalling pathways and metabotropic glutamate receptors, Brain Res. 869, 166–177, 2000. 130. Cooper, D.C. and White, F.J., L-type calcium channels modulate glutamatedriven bursting activity in the nucleus accumbens in vivo, Brain Res. 880, 212–218, 2000. 131. Stanley, E.F. and Atrakchi, A., Calcium currents recorded from a vertebrate presynaptic nerve terminal are resistant to the dihydropyridine nifedipine, Proc. Natl. Acad. Sci. USA 87, 9683–9687, 1990. 132. Qian, J. and Noebels, J.L., Presynaptic Ca2þ channels and neurotransmitter release at the terminal of a mouse cortical neuron, J. Neurosci. 21, 3721–3728, 2001. 133. McDonough, S.I., Boland, L.M., Mintz, I.M. and Bean, B.P., Interactions among toxins that inhibit N-type and P-type calcium channels, J. Gen. Physiol. 119, 313–328, 2002. 134. Mochida, S., Sheng, Z.H., Baker, C., Kobayashi, H. and Catterall, W.A., Inhibition of neurotransmission by peptides containing the synaptic protein interaction site of N-type Ca2þ channels, Neuron 17, 781–788, 1996. 135. Study, R.E., Isoflurane inhibits multiple voltage-gated calcium currents in hippocampal pyramidal neurons, Anesthesiology 81, 104–116, 1994. 136. Llina, R., Sugimori, M. and Simon, S.M., Transmission by presynaptic spike-like depolarization in the squid giant synapse, Proc. Natl. Acad. Sci. USA 79, 2415–2419, 1982. 137. Hall, A.C., Lieb, W.R. and Franks, N.P., Insensitivity of P-type calcium channels to inhalational and intravenous general anesthetics, Anesthesiology 81, 117–123, 1994. 138. Lynch, C., III and Pancrazio, J.J., Snails, spiders, and stereospecificity— is there a role for calcium channels in anesthetic mechanisms? Anesthesiology 81, 1–5, 1994. 139. Miao, N., Frazer, M.J. and Lynch, C., III, Volatile anesthetics depress Ca2þ transients and glutamate release in isolated cerebral synaptosomes, Anesthesiology 83, 593–603, 1995. 140. Larrabee, M.G. and Posternak, J.M., Selective action of anesthetics on synapses and axons in mammalian sympathetic ganglia, J. Neurophysiol. 15, 91–114, 1952. 141. Rehberg, B., Xiao, Y.-H. and Duch, D.S., Central nervous system sodium channels are significantly depressed at clinical concentrations of volatile anesthetics, Anesthesiology 84, 1223–1233, 1996. 142. Ratnakumari, L. and Hemmings, H.C., Inhibition of presynaptic sodium channels by halothane, Anesthesiology 88, 1043–1054, 1998. 143. Ratnakumari, L. and Hemmings, H.C., Effects of propofol on sodium channel-dependent sodium influx and glutamate release in rat cerebrocortical synaptosomes, Anesthesiology 86, 428–439, 1997. 144. Lingamaneni, R., Birch, M.L. and Hemmings, H.C., Widespread inhibition of sodium channel-dependent glutamate release from isolated nerve terminals by isoflurane and propofol, Anesthesiology 95, 1460–1466, 2001.
640
Stuth et al.
145. Wanner, S.G., Koch, R.O., Koschak, A., Trieb, M., Garcia, M.L., Kaczorowski, G.J. and Knaus, H.G., High-conductance calcium-activated potassium channels in rat brain: Pharmacology, distribution, and subunit composition, Biochemistry 38, 5392–5400, 1999. 146. Inanobe, A., Fujita, A., Ito, M., Tomoike, H., Inageda, K. and Kurachi, Y., Inward rectifier Kþ channel Kir2.3 is localized at the postsynaptic membrane of excitatory synapses, Am. J. Physiol. Cell. Physiol. 282, C1396– C1403, 2002. 147. Yost, C.S., Potassium channels, Anesthesiology 90, 1186–1203, 1999. 148. Rudy, B. and McBain, C.J., Kv3 channels: Voltage-gated Kþ channels designed for high-frequency repetitive firing, Trends Neurosci. 24, 517–526, 2001. 149. Bond, C.T., Maylie, J. and Adelman, J.P., Small-conductance calciumactivated potassium channels, Ann. NY Acad. Sci. 868, 370–378, 1999. 150. Brew, H.M. and Forsythe, I.D., Two voltage-dependent Kþ conductances with complementary functions in postsynaptic integration at a central auditory synapse, J. Neurosci. 15, 8011–8022, 1995. 151. Rudy, B., Chow, A., Lau, D., Amarillo, Y., Ozaita, A., Saganich, M., Moreno, H., Nadal, M.S., Hernandez-Pineda, R., Hernandez-Cruz, A., Erisir, A., Leonard, C. and Vega-Saenz de Miera, E., Contributions of Kv3 channels to neuronal excitability, Ann. NY Acad. Sci. 868, 304–343, 1999. 152. Browne, D.L., Gancher, S.T., Nutt, J.G., Brunt, E.R., Smith, E.A., Kramer, P. and Litt, M., Episodic ataxia/myokymia syndrome is associated with point mutations in the human potassium channel gene, KCNA1, Nat. Genet. 8, 136–140, 1994. 153. Kulkarni, R.S., Zorn, L.J., Anantharam, V., Bayley, H. and Treistman, S.N., Inhibitory effects of ketamine and halothane on recombinant potassium channels from mammalian brain, Anesthesiology 84, 900–909, 1999. 154. Haydon, D.A., Requena, J. and Simon, A.J., The potassium conductance of the resting squid axon and its blockage by clinical concentrations of general anesthetics, J. Physiol. (Lond.) 402, 363–374, 1988. 155. Elliott, J.R., Elliott, A.A., Harper, A.A. and Winpenny, J.P., Effects of general anaesthetics on neuronal sodium and potassium channels, Gen. Pharmacol. 23, 1005–1011, 1992. 156. Friederich, P. and Urban, B.W., Interaction of intravenous anesthetics with human neuronal potassium currents in relation to clinical concentrations, Anesthesiology 91, 1853–1860, 1999. 157. Dreixler, J.C., Jenkins, A., Cao, Y.J., Roizen, J.D. and Houamed, K.M., Patch-clamp analysis of anesthetic interactions with recombinant SK2 subtype neuronal calcium-activated potassium channels, Anesth. Analg. 90, 727–732, 2000. 158. Brown, A.M. and Birnbaumer, L., Ionic channels and their regulation by G protein subunits, Annu. Rev. Physiol. 52, 197–213, 1990. 159. North, R.A., Twelfth Gaddum memorial lecture. Drug receptors and the inhibition of nerve cells, Br. J. Pharmacol. 98, 13–28, 1989. 160. Gabriel, A., Abdallah, M., Yost, C.S., Winegar, B.D. and Kindler, C.H., Localization of the tandem pore domain Kþ channel KCNK5 (TASK-2) in the rat central nervous system, Brain Res. Mol. Brain Res. 98, 153–163, 2002.
Central Effects of General Anesthesia
641
161. Buckler, K.J., Background leak Kþ-currents and oxygen sensing in carotid body type 1 cells, Respir. Physiol. 115, 179–187, 1999. 162. Talley, E.M., Lei, Q., Sirois, J.E. and Bayliss, D.A., TASK-1, a two-pore domain K channel, is modulated by multiple neurotransmitters in motoneurons, Neuron 25, 399–410, 2000. 163. Franks, N.P. and Lieb, W.R., Background Kþ channels: An important target for volatile anesthetics? Nat. Neurosci. 2, 395–396, 1999. 164. Sirois, J.E., Lie, Q., Talley, E.M., Lynch, C., III and Bayliss, D.A., The TASK-1 two-pore domain Kþ channel is a molecular substrate for neuronal effects of inhalation anesthetics, J. Neurosci. 201, 6347–6354, 2000. 165. Nicoll, R.A. and Madison, D.V., General anesthetics hyperpolarize neurons in the vertebrate central nervous system, Science 217, 1055–1057, 1982. 166. Berg-Johnsen, J. and Langmoen, I.A., Isoflurane hyperpolarizes neurones in rat and human cerebral cortex, Acta Physiol. Scand. 130, 679–685, 1987. 167. Franks, N.P. and Lieb, W.R., Volatile general anaesthetics activate a novel Kþ current, Nature 333, 662–664, 1988. 168. Winegar, B.D., Owen, D.F., Yost, C.S., Forsayeth, J.R. and Mayeri, E., Volatile general anesthetics produce hyperpolarization of Aplysia neurons by activation of a discrete population of baseline potassium channels, Anesthesiology 85, 889–900, 1996. 169. Yamazaki, H., Haji, A., Okazaki, M. and Takeda, R., Immunoreactivity for glutamic acid decarboxylase and N-methyl-D-aspartate receptors of intracellularly labeled respiratory neurons in the cat, Neurosci. Lett. 293, 61–64, 2000. 170. Haji, A., Takeda, R. and Okazaki, M., Neuropharmacology of control of respiratory rhythm and pattern in mature mammals, Pharmacol. Ther. 86, 277– 304, 2000. 171. Robinson, D.M., Peebles, K.C., Kwok, H., Adams, B.M., Clarke, L.L., Woollard, G.A. and Funk, G.D., Prenatal nicotine exposure increases apnoea and reduces nicotinic potentiation of hypoglossal inspiratory output in mice, J. Physiol. (Lond.) 538, 957–973, 2002. 172. Franks, N.P. and Lieb, W.R., Molecular and cellular mechanisms of general anaesthesia, Nature 367, 607–614, 1994. 173. Yamakura, T., Bertaccini, E., Trudell, J.R. and Harris, R.A., Anesthetics and ion channels: Molecular models and sites of action, Annu. Rev. Pharmacol. Toxicol. 41, 23–51, 2001. 174. Richards, C.D. and White, A.E., The actions of volatile anaesthetics on synaptic transmission in the dentate gyrus, J. Physiol. (Lond.) 252, 241–257, 1975. 175. Richards, C.D. and Smaje, J.C., Anesthetics depress the sensitivity of cortical neurones to L-glutamate, Br. J. Pharmacol. 58, 347–357, 1976. 176. Mealing, G.A., Lanthorn, T.H., Murray, C.L., Small, D.L. and Morley, P., Differences in degree of trapping of low-affinity uncompetitive N-methyl-Daspartic acid receptor antagonists with similar kinetics of block, J. Pharmacol. Exp. Ther. 288, 204–210, 1999. 177. Orser, B.A., Pennefather, P.S. and MacDonald, J.F., Multiple mechanisms of ketamine blockade of N-methyl-D-aspartate receptors, Anesthesiology 86, 903–917, 1997.
642
Stuth et al.
178. MacDonald, J.F., Bartlett, M.C., Mody, I., Pahapill, P., Reynolds, J.N., Salter, M.W., Schneiderman, J.H. and Pennefather, P.S., Actions of ketamine, phencyclidine and MK-801 on NMDA receptor currents in cultured mouse hippocampal neurones, J. Physiol. (Lond.) 432, 483–508, 1991. 179. Anis, N.A., Berry, S.C., Burton, N.R. and Lodge, D., The dissociative anaesthetics, ketamine and phencyclidine, selectively reduce excitation of central mammalian neurones by N-methyl-D-aspartate, Br. J. Pharmacol. 79, 565–575, 1983. 180. Cheng, G. and Kendig, J.J., Enflurane directly depresses glutamate AMPA and NMDA currents in mouse spinal cord motor neurons independent of actions on GABAA or glycine receptors, Anesthesiology 93, 1075–1084, 2000. 181. Kirson, E.D., Yaari, Y. and Perouansky, M., Presynaptic and postsynaptic actions of halothane at glutamatergic synapses in the mouse hippocampus, Br. J. Pharmacol. 124, 1607–1614, 1998. 182. de Sousa, S.L.M., Dickinson, R., Lieb, W.R. and Franks, N.P., Contrasting synaptic actions of the inhalational general anesthetics isoflurane and xenon, Anesthesiology 92, 1055–1066, 2000. 183. Ming, Z., Griffith, B.L., Breese, G.R., Mueller, R.A. and Criswell, H.E., Changes in the effect of isoflurane on N-methyl-D-aspartic acid-gated currents in cultured cerebral cortical neurons with time in culture: Evidence for subunit specificity, Anesthesiology 97, 856–867, 2002. 184. Orser, B.A., Bertlik, M., Wang, L.Y. and MacDonald, J.F., Inhibition by propofol (2,6 di-isopropylphenol) of the N-methyl-D-aspartate subtype of glutamate receptor in cultured hippocampal neurones, Br. J. Pharmacol. 116, 1761–1768, 1995. 185. Yamakura, T., Sakimura, K., Shimoji, K. and Mishina, M., Effects of propofol on various AMPA-, kainate- and NMDA-selective glutamate receptor channels expressed in Xenopus oocytes, Neurosci. Lett. 188, 187–190, 1995. 186. Franks, N.P., Dickinson, R., de Sousa, S.L., Hall, A.C. and Lieb, W.R., How does xenon produce anesthesia? Nature 396, 324, 1998. 187. Yamakura, T. and Harris, R.A., Effects of gaseous anesthetics nitrous oxide and xenon on ligand-gated ion channels: Comparison with isoflurane and ethanol, Anesthesiology 93, 1095–1101, 2000. 188. Peoples, R.W. and Weight, F.F., Anesthetic actions on excitatory amino acid receptors, in Anesthesia: Biologic Foundations, Yaksh, T.L., Lynch, C., III, Zapol, W.M., Maze, M., Biebuyck, J.F. and Saidman, L.J., eds., Philadelphia, Lippincott-Raven Publishers, pp. 239–258, 1997. 189. Evers, A.S. and Steinbach, J.H., Supersensitive sites in the central nervous system, Anesthesiology 86, 760–762, 1997. 190. Ballantyne, D. and Scheid, P., Central respiratory chemosensitivity: Cellular and network mechanisms, Adv. Exp. Med. Biol. 499, 17–26, 2001. 191. Alcayaga, J., Iturriaga, R., Varas, R., Arroyo, J. and Zapata, P., Selective activation of carotid nerve fibers by acetylcholine applied to the cat petrosal ganglion in vitro, Brain Res. 786, 47–54, 1998. 192. Haxhiu, M.A., Mitra, J., van Lunteren, E., Bruce, E.N. and Cherniack, N.S., Hypoglossal and phrenic responses to cholinergic agents applied to ventral medullary surface, Am. J. Physiol. 247, R939–R944, 1984.
Central Effects of General Anesthesia
643
193. Haxhiu, M.A., Mitra, J., van Lunteren, E., Prabhakar, N.R. and Cherniack, N.S., Influence of central chemoreceptor afferent inputs on respiratory muscle activity, Am. J. Physiol. 249, R266–R273, 1985. 194. Takeda, R. and Haji, A., Synaptic response of bulbar respiratory neurons in hypercapnic stimulation in peripherally chemodenervated cats, Brain Res. 561, 307–317, 1991. 195. Andoh, T., Furuya, R., Oka, K., Hattori, S., Watanabe, I., Kamiya, Y. and Okumura, F., Differential effects of thiopental on neuronal nicotinic acetylcholine receptors and P2X purinergic receptors in PC12 cells, Anesthesiology 87, 1199–1209, 1997. 196. Flood, P., Ramirez-Latorre, J. and Role, L., Alpha 4 beta 2 neuronal nicotinic acetylcholine receptors in the central nervous system are inhibited by isoflurane and propofol, but alpha7-type nicotinic acetylcholine receptors are unaffected, Anesthesiology 86, 859–865, 1997. 197. Violet, J.M., Downie, D.L., Nakisa, R.C., Lieb, W.R. and Franks, N.P., Differential sensitivities of mammalian neuronal and muscle nicotinic acetylcholine receptors to general anesthetics, Anesthesiology 86, 866–874, 1997. 198. Borghese, C.M. and Harris, R.A., Anesthetic-induced immobility: Neuronal nicotinic acetylcholine receptors are no longer in the picture, Anesth. Analg. 95, 509–511, 2002. 199. Raines, D.E., Claycomb, R.J. and Forman, S.A., Nonhalogenated anesthetic alkanes and perhalogenated nonimmobilizing alkanes inhibit alpha 4 beta 2 neuronal nicotinic acetylcholine receptors, Anesth. Analg. 94, 573–577, 2002. 200. Raines, D.E., Claycomb, R.J., Scheller, M. and Forman, S.A., Nonhalogenated alkane anesthetics fail to potentiate agonist actions on two ligand-gated ion channels, Anesthesiology 95, 470–477, 2001. 201. Okazaki, M., Takeda, R., Haji, A. and Yamazaki, H., Glutamic acid decarboxylase-immunoreactivity of bulbar respiratory neurons identified by intracellular recording and labeling in rats, Brain Res. 914, 34–47, 2001. 202. Banks, M.I. and Pearce, R.A., Dual actions of volatile anesthetics on GABAA IPSCs: Dissociation of blocking and prolonging effects, Anesthesiology 90, 120–134, 1999. 203. Tanelian, D.L., Kosek, P., Mody, I. and MacIver, M.B., The role of the GABAA receptor/chloride channel complex in anesthesia, Anesthesiology 78, 757–776, 1993. 204. MacIver, M.B., General anesthetic actions on transmission at glutamate and GABA synapses, in Anesthesia: Biologic Foundations, Yaksh, T.L., Lynch, C., III, Zapol, W.M., Maze, M., Biebuyck, J.F. and Saidman, L.J., eds., Philadelphia, Lippincott-Raven Publishers, pp. 277–286, 1997. 205. Lin, L.-H., Chen, L.L., Zirrolli, J.A. and Harris, R.A., General anesthetics potentiate g-aminobutyric acid actions on g-aminobutyric acidA receptors expressed by Xenopus oocytes: Lack of involvement of intracellular calcium, J. Pharmacol. Exp. Ther. 263, 569–578, 1992. 206. Jones, M.V. and Harrison, N.L., Effects of volatile anesthetics on the kinetics of inhibitory postsynaptic currents in cultured rat hippocampal neurons, J. Neurophysiol. 70, 1339–1349, 1993.
644
Stuth et al.
207. Jones, M.V., Brooks, P.A. and Harrison, N., Enhancement of g-aminobutyric acid-activated Cl currents in cultured rat hippocampal neurones by three volatile anaesthetics, J. Physiol. (Lond.) 449, 279–293, 1992. 208. Banks, M.I. and Pearce, R.A., Kinetic differences between synaptic and extrasynaptic GABAA receptors in CA1 pyramidal cells, J. Neurophysiol. 20, 937–948, 2000. 209. Brickley, S.G., Cull-Candy, S.G. and Farrant, M., Development of a tonic form of synaptic inhibition in rat cerebellar granule cells resulting from persistent activation of GABAA receptors, J. Physiol. (Lond.) 497, 753–759, 1996. 210. Moody, E.J., Harris, B.D. and Skolnick, P., Stereospecific actions of the inhalation anesthetic isoflurane at the GABAA receptor complex, Brain Res. 615, 101–106, 1993. 211. Harris, B., Moody, E. and Skolnick, P., Isoflurane anesthesia is stereoselective, Eur. J. Pharmacol. 217, 215–216, 1992. 212. Chery, N. and de Koninck, Y., Junctional versus extrajunctional glycine and GABAA receptor-mediated IPSCs in identified lamina I neurons of the adult rat spinal cord, J. Neurosci. 19, 7342–355, 1999. 213. Siebler, M., Pekel, M., Koller, H. and Muller, H.W., Strychnine-sensitive glycine receptors in cultured primary neurons from rat neocortex, Brain Res. Dev. Brain Res. 2, 289–292, 1973. 214. Schmid, K., Foutz, A.S. and Denavit-Saubie´, M., Inhibitions mediated by glycine and GABAA receptors shape the discharge patterns of bulbar respiratory neurons, Brain Res. 710, 150–160, 1996. 215. Downie, D.L., Hall, A.C., Lieb, W.R. and Franks, N.P., Effects of inhalational anaesthetics on native glycine receptors in rat medullary neurones and recombinant glycine receptors in Xenopus oocytes, Br. J. Pharmacol. 118, 493–502, 1996. 216. Mihic, S.J., Ye, Q., Wick, M.J., Koltchine, V.V., Krasowski, M.D., Finn, S.E., Mascia, M.P., Valenzuela, C.F., Hanson, K.K., Greenblatt, E.P., Harris, R.A. and Harrison, N.L., Sites of alcohol and volatile anaesthetic action on GABA sub A and glycine receptors, Nature 389, 385–389, 1997. 217. Mascia, M.P., Machu, T.K. and Harris, R.A., Enhancement of homomeric glycine receptor function by long-chain alcohols and anaesthetics, Br. J. Pharmacol. 119, 1331–1336, 1996. 218. Yamakura, T., Chavez-Noriega, L.E. and Harris, R.A., Subunit-dependent inhibition of human neuronal nicotinic acetylcholine receptors and other ligand-gated ion channels by dissociative anesthetics ketamine and dizocilpine, Anesthesiology 92, 1144–1153, 2000. 219. Hara, M., Kai, Y. and Ikemoto, Y., Enhancement by propofol of the g-aminobutyric acidA response in dissociated hippocampal pyramidal neurons of the rat, Anesthesiology 81, 988–994, 1994. 220. Tryfon, S., Kontakiotis, T., Mavrofridis, E. and Patakas, D., HeringBreuer reflex in normal adults and in patients with chronic obstructive pulmonary disease and interstitial fibrosis, Respiration 68, 140–144, 2001. 221. Tassonyi, E., Charpantier, E., Muller, D., Dumont, L. and Bertrand, D., The role of nicotinic acetylcholine receptors in the mechanisms of anesthesia, Brain Res. 57, 133–150, 2002.
Central Effects of General Anesthesia
645
222. Feldman, J.L., Neurophysiology of breathing in mammals, in Handbook of Physiology, The Nervous System, Brown, F.E., ed., 1st edn., Bethesda, MD, American Physiological Society, pp. 463–523, 1986. 223. von Euler, C., Brain stem mechanisms for generation and control of breathing pattern, in Handbook of Physiology, The Respiratory System, Cherniack, N.S. and Widdecombe, J.G., eds., 3rd edn., Bethesda, MD, American Physiological Society, pp. 1–67, 1986. 224. Ezure, K., Synaptic connections between medullary respiratory neurons and considerations on the genesis of respiratory rhythm, Prog. Neurobiol. 35, 429–450, 1990. 225. St. John, W.M., Neurogenesis of patterns of automatic ventilatory activity, Prog. Neurobiol. 56, 97–117, 1998. 226. Bianchi, A.L., Denavit-Saubie´, M. and Champagnat, J., Central control of breathing in mammals: Neuronal circuitry, membrane properties, and neurotransmitters, Physiol. Rev. 75, 1–45, 1995. 227. Richter, D.W., Generation and maintenance of the respiratory rhythm, J. Exp. Biol. 100, 93–108, 1982. 228. Ramirez, J.M. and Richter, D.W., The neuronal mechanisms of respiratory rhythm generation, Curr. Opin. Neurobiol. 6, 817–825, 1996. 229. Richter, D.W. and Spyer, K.M., Studying rhythmogenesis of breathing: Comparison of in vivo and in vitro models, Trends Neurosci. 24, 464–472, 2001. 230. Feldman, J.L. and McCrimmon, D.R., Neural control of breathing, in Fundamental Neuroscience, Bloom, F.E., Landis, S.C., Roberts, J.L., Squire, L.C. and Zigmond, M.J., eds., New York, Academic Press, pp. 1063–1090, 1998. 231. Kirkwood, P.A., Munson, J.B. and Sears, T.A., Respiratory interneurones in the thoracic spinal cord of the cat, J. Physiol. (Lond.) 395, 161–192, 1988. 232. Davies, J.G.M., Kirkwood, P.A. and Sears, T.A., The detection of monosynaptic connexions from inspiratory bulbospinal neurones to inspiratory motoneurones in the cat, J. Physiol. (Lond.) 368, 33–62, 1985. 233. Davies, J.G.M., Kirkwood, P.A. and Sears, T.A., The distribution of monosynaptic connexions from inspiratory bulbospinal neurones to inspiratory motoneurones in the cat, J. Physiol. (Lond.) 368, 63–87, 1985. 234. Lipski, J. and Duffin, J., An electrophysiological investigation of propriospinal inspiratory neurons in the upper cervical cord of the cat, Exp. Brain. Res. 61, 625–637, 1986. 235. Sears, T.A., Central rhythm generation and spinal integration, Chest 97, 45S–51S, 1990. 236. Smith, J.C., Ellenberger, H.H., Ballanyi, K., Richter, D.W. and Feldman, J.L., Pre-Bo¨tzinger complex: A brainstem region that may generate respiratory rhythm in mammals, Science 254, 726–729, 1991. 237. Johnson, S.M., Smith, J.C., Funk, G.D. and Feldman, J.L., Pacemaker behavior of respiratory neurons in medullary slices from neonatal rat, J. Neurophysiol. 72, 2598–2608, 1994. 238. Koshiya, N. and Smith, J.C., Neuronal pacemaker for breathing visualized in vitro, Nature 400, 360–363, 1999.
646
Stuth et al.
239. Thoby-Brisson, M. and Ramirez, J.M., Role of inspiratory pacemaker neurons in mediating the hypoxic response of the respiratory network in vitro, J. Neurosci. 20, 5858–5866, 2000. 240. Thoby-Brisson, M. and Ramirez, J.M., Identification of two types of inspiratory pacemaker neurons in the isolated respiratory neural network of mice, J. Neurophysiol. 86, 104–112, 2001. 241. Zuperku, E.J., Hopp, F.A. and Kampine, J.P., Respiratory oscillator model based on functional correlates derived from pulmonary reflexes, Physiologist 25, 188, 1982. 242. Balis, U.J., Morris, K.F., Koleski, J. and Lindsey, B.G., Simulations of a ventrolateral medullary neural network for respiratory rhythmogenesis inferred from spike train cross-corelation, Biol. Cybern. 70, 311–327, 1994. 243. Ogilvie, M.D., Gottschalk, A., Anders, K., Richter, D.W. and Pack, A.I., A network model of respiratory rhythmogenesis, Am. J. Physiol. 263, R962–R975, 1992. 244. Duffin, J., Ezure, K. and Lipski, J., Breathing rhythm generation: Focus on the rostral ventrolateral medulla, NIPS 10, 133–140, 1995. 245. Rybak, I.A., Paton, J.F.R. and Schwaber, J.S., Modeling neural mechanisms for genesis of respiratory rhythm and pattern. II. Network models of the central respiratory neurons, J. Neurophysiol. 77, 2007–2026, 1997. 246. Paton, J.F.R., Ramirez, J.M. and Richter, D.W., Mechanisms of respiratory rhythm generation change profoundly during early life in mice and rats, Neurosci. Lett. 170, 167–170, 1994. 247. Clark, F.J. and von Euler, C., On the regulation of depth and rate of breathing, J. Physiol. (Lond.) 222, 267–295, 1972. 248. Zuperku, E.J. and Hopp, F.A., On the relation between expiratory duration and subsequent inspiratory duration, J. Appl. Physiol. 58, 419–430, 1985. 249. Hopp, F.A., Seagard, J.L., Bajic, J. and Zuperku, E.J., Respiratory responses to aortic and carotid chemoreceptor activation in the dog, J. Appl. Physiol. 70, 2539–2550, 1991. 250. Berkenbosch, A., van Beek, J.H.G.M., Olievier, C.N., De Goede, J. and Quanjer, P.H., Central respiratory CO2 sensitivity at extreme hypocapnia, Respir. Physiol. 55, 95–102, 1984. 251. Haji, A., Remmers, J.E., Connelly, C. and Takeda, R., Effects of glycine and GABA on bulbar respiratory neurons of cat, J. Neurophysiol. 63, 955–965, 1990. 252. Haji, A., Takeda, R. and Remmers, J.E., Evidence that glycine and GABA mediate postsynaptic inhibition of bulbar respiratory neurons in the cat, J. Appl. Physiol. 73, 2333–2342, 1992. 253. Cohen, M.I., Discharge patterns of brain-stem respiratory neurons in relation to carbon dioxide tension, J. Neurophysiol. 31, 142–165, 1968. 254. St. John, W.M. and Bianchi, A.L., Responses of bulbospinal and laryngeal respiratory neurons to hypercapnia and hypoxia, J. Appl. Physiol. 59, 1201–1207, 1985. 255. Lumb, A.B., Control of breathing, in Nunn’s Applied Respiratory Physiology, 5th edn., Oxford, Butterworth-Heinemann, pp. 82–112, 2000.
Central Effects of General Anesthesia
647
256. Bainton, C.R., Kirkwood, P.A. and Sears, T.A., On the transmission of the stimulating effects of carbon dioxide to the muscles of respiration, J. Physiol. (Lond.) 280, 249–272, 1978. 257. Bainton, C.R. and Kirkwood, P.A., The effect of carbon dioxide on the tonic and the rhythmic discharges of expiratory bulbospinal neurones, J. Physiol. (Lond.) 296, 291–314, 1979. 258. Sears, T.A., Berger, A.J. and Phillipson, E.A., Reciprocal tonic activation of inspiratory and expiratory motoneurones by chemical drives, Nature 299, 728–730, 1982. 259. Fink, B.R., Influence of cerebral activity in wakefulness on regulation of breathing, J. Appl. Physiol. 16, 15–20, 1961. 260. Orem, J., The nature of the wakefulness stimulus for breathing, Prog. Clin. Biol. Res. 345, 23–30, 1990. 261. Heeringa, J., Berkenbosch, A., DeGoede, J. and Olievier, C.N., Relative contribution of central and peripheral chemoreceptors to the ventilatory response to CO2 during hyperoxia, Respir. Physiol. 37, 365–379, 1979. 262. Smith, C.A., Jameson, L.C., Mitchell, G.S., Musch, T.I. and Dempsey, J.A., Central-peripheral chemoreceptor interaction in awake cerebrospinal fluidperfused goats, J. Appl. Physiol. 56, 1541–1549, 1984. 263. Mitchell, R.A., Loeschcke, H.H., Massion, W.H. and Severinghaus, J.W., Respiratory responses mediated through superficial chemosensitive areas on the medulla, J. Appl. Physiol. 18, 523–533, 1963. 264. Loeschcke, H.H., Central chemosensitivity and the reaction theory, J. Physiol. (Lond.) 332, 1–24, 1982. 265. Hayashi, F. and Lipski, J., The role of inhibitory amino acids in control of respiratory motor output in an arterially perfused rat, Respir. Physiol. 89, 47–63, 1992. 266. Ballanyi, K., Onimaru, H. and Homma, I., Respiratory network function in the isolated brainstem-spinal cord of newborn rats, Prog. Neurobiol. 59, 583–634, 1999. 267. Bajic, J., Zuperku, E.J., Tonkovic-Capin, M. and Hopp, F.A., Interaction between chemoreceptor and stretch receptor inputs at medullary respiratory neurons, Am. J. Physiol. 266, R1951–R1961, 1994. 268. Tonkovic-Capin, M., Zuperku, E.J., Stuth, E.A., Bajic, J., Dogas, Z. and Hopp, F.A., Effect of central CO2 drive on lung inflation responses of expiratory bulbospinal neurons in dogs, Am. J. Physiol. 279, R1606–R1618, 2000. 269. Bajic, J., Zuperku, E.J., Tonkovic-Capin, M. and Hopp, F.A., Expiratory bulbospinal neurons of dogs: I. Control of discharge patterns by pulmonary stretch receptors, Am. J. Physiol. 262, R1075–R1086, 1992. 270. Iscoe, S., Control of abdominal muscles, Prog. Neurobiol. 56, 433–506, 1998. 271. Richter, D.W., Camerer, H., Meesmann, M. and Rohrig, N., Studies on the synaptic interconnection between bulbar respiratory neurons of cats, Pflu¨gers Arch. 380, 245–257, 1979. 272. Richter, D.W., Ballantyne, D. and Remmers, J.E., The differential organization of medullary post-inspiratory activities, Pflu¨gers Arch. 410, 420–427, 1987.
648
Stuth et al.
273. Ballantyne, D. and Richter, D.W., Post-synaptic inhibition of bulbar inspiratory neurones in the cat, J. Physiol. (Lond.) 384, 67–87, 1984. 274. Ballantyne, D. and Richter, D.W., The non-uniform character of expiratory synaptic activity in expiratory bulbospinal neurones of the cat, J. Physiol. (Lond.) 370, 433–456, 1986. 275. Bianchi, A.L., Grelot, L., Iscoe, S. and Remmers, J.E., Electrophysiological properties of rostral medullary respiratory neurones in the cat: An intracellular study, J. Physiol. (Lond.) 407, 293–310, 1988. 276. Takeda, R. and Haji, A., Mechanisms underlying post-inspiratory depolarization in post-inspiratory neurons of the cat, Neurosci. Lett. 150, 1–4, 1993. 277. Pierrefiche, O., Schmid, K., Foutz, A.S. and Denavit-Saubie´, M., Endogenous activation of NMDA and non-NMDA glutamate receptors on respiratory neurones in cat medulla, Neuropharmacology 30, 429–440, 1991. 278. Pierrefiche, O., Foutz, A.S., Champagnat, J. and Denavit-Saubie´, M., The bulbar network of respiratory neurons during apneusis induced by a blockade of NMDA receptors, Exp. Brain. Res. 89, 623–639, 1992. 279. Connelly, C.A., Otto-Smith, M.R. and Feldman, J.L., Blockade of NMDA receptor-channels by MK-801 alters breathing in adult rats, Brain Res. 596, 99–110, 1992. 280. Dogas, Z., Stuth, E.A.E., Hopp, F.A., McCrimmon, D.R. and Zuperku, E.J., NMDA receptor-mediated transmission of carotid body chemoreceptor input to expiratory bulbospinal neurones in dogs, J. Physiol. (Lond.) 487, 639–651, 1995. 281. Dogas, Z., In vivo activity of canine expiratory bulbospinal neurons is differentially affected by two non-N-methyl-D-aspartate receptor antagonists, Croat. Med. J. 37, 15–20, 1996. 282. Krolo, M., Stuth, E.A., Tonkovic-Capin, M., Dogas, Z., Hopp, F.A., McCrimmon, D.R., Zuperku, E.J., Differential roles of ionotropic glutamate receptors in canine medullary inspiratory neurons of the ventral respiratory group, J. Neurophysiol. 82, 60–68, 1999. 283. Gozal, D., Potentiation of hypoxic ventilatory response by hyperoxia in the conscious rat: Putative role of nitric oxide, J. Appl. Physiol. 85, 129–132, 1998. 284. Stucke, A.G., Stuth, E.A.E., Tonkovic-Capin, V., Kampine, J.P. and Zuperku, E.J., Effects of halothane on excitatory neurotransmission to medullary inspiratory neurons in a decerebrate dog model, Anesthesiology A1337, 2002. 285. Krolo, M., Stuth, E.A., Tonkovic-Capin, M., Hopp, F.A., McCrimmon, D.R. and Zuperku, E.J., Relative magnitude of tonic and phasic synaptic excitation of medullary inspiratory neurons in dogs, Am. J. Physiol. 279, R639–R649, 2000. 286. Foutz, A.S., Champagnat, J. and Denavit-Saubie´, M., Involvement of N-methyl-D-aspartate (NMDA) receptors in respiratory rhythmogenesis, Brain Res. 500, 199–208, 1989. 287. Ling, L., Karius, D.R. and Speck, D.F., Role of N-methyl-D-aspartate receptors in the pontine pneumotaxic mechanism in the cat, J. Appl. Physiol. 76, 1138–1143, 1994.
Central Effects of General Anesthesia
649
288. Karius, D.R., Ling, L. and Speck, D.F., Nucleus tractus solitarius and excitatory amino acids in efferent-evoked inspiratory termination, J. Appl. Physiol. 76, 1293–1301, 1994. 289. Berger, I., Gillis, R.A., Vitagliano, S., Panico, W.H., Magee, S., Kelly, M., Norman, W.P., McManigle, J.E. and Taveira DaSilva, A.M., NMDA receptors are involved at the ventrolateral nucleus tractus solitarii for termination of inspiration, Eur. J. Pharmacol. 277, 195–208, 1995. 290. Haji, A., Okazaki, M., Yamazaki, H. and Takeda, R., NMDA receptormediated inspiratory off-switching in pneumotaxic-disconnected cats, Neurosci. Res. 32, 323–331, 1998. 291. Champagnat, J., Denavit-Saubie´, M., Moyanova, S. and Rondouin, G., Involvement of amino acids in periodic inhibitions of bulbar respiratory neurons, Brain Res. 237, 351–365, 1982. 292. Dogas, Z., Krolo, M., Stuth, E.A., Tonkovic-Capin, M., Hopp, F.A., McCrimmon, D.R. and Zuperku, E.J., Differential effects of GABAA receptor antagonists in the control of respiratory neuronal discharge patterns, J. Neurophysiol. 80, 2368–2377, 1998. 293. Kawai, A., Ballantyne, D., Muckenhoff, K. and Scheid, P., Chemosensitive medullary neurones in the brainstem-spinal cord preparation of the neonatal rat, J. Physiol. (Lond.) 492, 277–292, 1996. 294. Haji, A., Okazaki, M. and Takeda, R., GABAA receptor-mediated inspiratory termination evoked by vagal stimulation in decerebrate cats, Neuropharmacology 38, 1261–1272, 1999. 295. McCrimmon, D.R., Zuperku, E.J., Hayashi, F., Dogas, Z., Hinrichsen, C.F., Stuth, E.A., Tonkovic-Capin, M., Krolo, M. and Hopp, F.A., Modulation of the synaptic drive to respiratory premotor and motor neurons, Respir. Physiol. 110, 161–176, 1997. 296. Zuperku, E.J. and McCrimmon, D.R., Gain modulation of respiratory neurons, Respir. Physiol. Neurobiol. 131, 121–133, 2002. 297. McCrimmon, D.R., Mitchell, G.S. and Dekin, M.S., Glutamate, GABA, and serotonin in ventilatory control, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., 2nd edn., New York, Marcel Dekker, Inc., pp. 151– 218, 1995. 298. Rekling, J.C., Funk, G.D., Bayliss, D.A., Dong, X.W. and Feldman, J.L., Synaptic control of motoneuronal excitability, Physiol. Rev. 80, 767–852, 2000. 299. Feldman, J.L. and Smith, J.C., Neural control of respiratory pattern in mammals: An overview, in Regulation of Breathing, Dempsey, J.A. and Pack, A.I., eds., 2nd edn., New York, Marcel Dekker, Inc., pp. 39–68, 1995. 300. Lalley, P.M., Effects of baclofen and gamma-aminobutyric acid on different types of medullary respiratory neurons, Brain Res. 376, 392–395, 1986. 301. Haji, A. and Takeda, R., Microiontophoresis of baclofen on membrane potential and input resistance in bulbar respiratory neurons in the cat, Brain Res. 622, 294–298, 1993. 302. Pierrefiche, O., Foutz, A.S. and Denavit-Saubie´, M., Effects of GABAB receptor agonists and antagonists on the bulbar respiratory network in cat, Brain Res. 605, 77–84, 1993.
650
Stuth et al.
303. Zuperku, E.J., Coon, R.L., Bosnjak, Z.J., Igler, F.O. and Kampine, J.P., Pulmonary effects of halothane on the breathing pattern, Proc. Am. Soc. Anesth. 209–210, 1976. 304. Miserocchi, G. and Sant’Ambrogio, G., Responses of pulmonary stretch receptors to static pressure inflations, Respir. Physiol. 21, 77–85, 1974. 305. Iscoe, S., Pulmonary stretch receptor discharge patterns in eupnea, hypercapnia, and hypoxia, J. Appl. Physiol. 53, 346–354, 1982. 306. Nilsestuen, J.O., Coon, R.L., Woods, M. and Kampine, J.P., Location of lung receptors mediating the breathing frequency response to pulmonary CO2, Respir. Physiol. 45, 343–355, 1981. 307. Miserocchi, G. and Sant’Ambrogio, G., Distribution of pulmonary stretch receptors in the intrapulmonary airways of the dog, Respir. Physiol. 21, 71–75, 1974. 308. Bartoli, A., Bystrzycka, E., Guz, A., Jain, S.K. and Noble, M.I.M., Studies of the pulmonary vagal control of central respiratory rhythm in the absence of breathing movements, J. Physiol. (Lond.) 230, 449–465, 1973. 309. Zuperku, E.J., Coon, R.L., Brill, J.J. and Kampine, J.P., The effects of halothane, ethrane, and methoxyfluorane on pulmonary stretch receptor afferent nerve activity in the dog, Proc. ASA Ann. Mtg. 181–182, 1972. 310. Coleridge, H.M., Coleridge, J.C.G., Luck, J.C. and Norman, J., The effect of four volatile anaesthetic agents on the impulse activity of two types of pulmonary receptor, Br. J. Anaesth. 40, 484–492, 1968. 311. Knox, C.K., Characteristics of inflation and deflation reflexes during expiration in the cat, J. Neurophysiol. 36, 284–295, 1973. 312. Zuperku, E.J., Hopp, F.A. and Kampine, J.P., Central integration of pulmonary stretch receptor input in the control of expiration, J. Appl. Physiol. 52, 1296–1315, 1982. 313. Nilsestuen, J.O., Coon, R.L., Igler, F.O., Zuperku, E.J. and Kampine, J.P., Breathing frequency responses to pulmonary CO2 in an isolated lobe of the canine lung, J. Appl. Physiol. 47, 1201–1206, 1979. 314. Gaudy, J.H., The Hering-Breuer reflex in man? Br. J. Anaesth. 66, 627–628, 1991. 315. Rabbette, P.S., Fletcher, M.E., Dezateux, C.A., Soriano-Brucher, H. and Stocks, J., Hering-Breuer reflex and respiratory system compliance in the first year of life: A longitudinal study, J. Appl. Physiol. 76, 650–656, 1994. 316. Kochi, T., Izumi, Y., Isono, S., Ide, T. and Mizuguchi, T., Breathing pattern and occlusion pressure waveform in humans anesthetized with halothane or sevoflurane, Anesth. Analg. 73, 327–332, 1991. 317. Stuth, E.A.E., Dogas, Z., Krolo, M., Kampine, J.P., Hopp, F.A. and Zuperku, E.J., Dose-dependent effects of halothane on the phrenic nerve responses to acute hypoxia in vagotomized dogs, Anesthesiology 87, 1428–1439, 1997. 318. Stuth, E.A.E., Dogas, Z., Krolo, M., Kampine, J.P., Hopp, F.A. and Zuperku, E.J., Dose-dependent effects of halothane on the phrenic nerve responses to carbon dioxide mediated by the carotid body chemoreceptors in vagotomized dogs, Anesthesiology 87, 1440–1449, 1997.
Central Effects of General Anesthesia
651
319. Tonkovic-Capin, V., Stucke, A.G., Stuth, E.A., Tonkovic-Capin, M., Krolo, M., Hopp, F.A., McCrimmon, D.R. and Zuperku, E.J., Differential modulation of respiratory neuronal discharge patterns by GABAA receptor and apamin-sensitive Kþ channel antagonism, J. Neurophysiol. 86, 2363–2373, 2001. 320. Tonkovic-Capin, M., Krolo, M., Stuth, E.A.E., Hopp, F.A. and Zuperku, E.J., Improved method of canine decerebration, J. Appl. Physiol. 85, 747–750, 1998. 321. Tonkovic-Capin, V., Stucke, A.G., Stuth, E.A., Tonkovic-Capin, M., Hopp, F.A., McCrimmon, D.R. and Zuperku, E.J., Differential processing of excitation by GABAergic gain modulation in canine caudal ventral respiratory group neurons, J. Neurophysiol. 89, 862–870, 2003. 322. Perouansky, M., Baranov, D., Salman, M. and Yaari, Y., Effects of halothane on glutamate receptor-mediated excitatory postsynaptic currents, Anesthesiology 83, 109–119, 1995. 323. Haji, A., Pierrefiche, O., Takeda, R., Foutz, A.S., Champagnat, J. and Denavit-Saubie´, M., Membrane potentials of respiratory neurones during dizocilpine-induced apneusis in adult cats, J. Physiol. (Lond.) 495, 851–861, 1996. 324. Nishikawa, K. and MacIver, M.B., Membrane and synaptic actions of halothane on rat hippocampal pyramidal neurons and inhibitory interneurons, J. Neurosci. 20, 5915–5923, 2000. 325. Remmers, J.E., deGroot, W.J., Sauerland, E.K. and Anch, A.M., Pathogenesis of upper airway occlusion during sleep, J. Appl. Physiol. 44, 931–938, 1978. 326. Viana, F., Bayliss, D.A. and Berger, A.J., Repetitive firing properties of developing rat brainstem motoneurones, J. Physiol. (Lond.) 486, 745–761, 1995. 327. Bayliss, D.A., Viana, F., Talley, E.M. and Berger, A.J., Neuromodulation of hypoglossal motoneurons: Cellular and developmental mechanisms, Respir. Physiol. 110, 139–150, 1997. 328. Berger, A.J., Determinants of respiratory motoneuron output, Respir. Physiol. 122, 259–269, 2000. 329. O’Brien, J.A., Isaacson, J.S. and Berger, A.J., NMDA and non-NMDA receptors are co-localized at excitatory synapses of rat hypoglossal motoneurons, Neurosci. Lett. 227, 5–8, 1997. 330. O’Brien, J.A. and Berger, A.J., Cotransmission of GABA and glycine to brain stem motoneurons, J. Neurophysiol. 82, 1638–1641, 1999. 331. Withington-Wray, D.J., Mifflin, S.W. and Spyer, K.M., Intracellular analysis of respiratory-modulated hypoglossal motoneurons in the cat, Neuroscience 25, 1041–1051, 1988. 332. Qian, N. and Sejnowski, T.J., When is an inhibitory synapse effective? Proc. Natl. Acad. Sci. USA 87, 8145–8149, 1990. 333. Bayliss, D.A., Talley, E.M., Sirois, J.E. and Lei, Q., TASK-1 is a highly modulated pH-sensitive ‘leak’ Kþ channel expressed in brainstem respiratory neurons, Respir. Physiol. 129, 159–174, 2001. 334. Washburn, C.P., Sirois, J.E., Talley, E.M., Guyenet, P.G. and Bayliss, D.A., Serotonergic raphe neurons express TASK channel transcripts and a
652
335.
336.
337.
338.
339.
340.
341.
342. 343.
344.
345.
346. 347.
Stuth et al. TASK-like pH- and halothane-sensitive Kþ conductance, J. Neurosci. 22, 1256–1265, 2002. Woch, G. and Kubin, L., Non-reciprocal control of rhythmic activity in respiratory-modulated XII motoneurons, Neuroreport 6, 2085– 2088, 1995. Hickey, R.F., Fourcade, H.E., Eger, E.I., 2d, Larson, C.P., Jr., Bahlman, S.H., Stevens, W.C., Gregory, G.A. and Smith, N.T., The effects of ether, halothane, and forane on apneic thresholds in man, Anesthesiology 35, 32–37, 1971. Franks, N.P. and Lieb, W.R., Temperature dependence of the potency of volatile anesthetics: Implications for in vitro experiments, Anesthesiology 84, 716–720, 1996. Mikulec, A.A., Pittson, S., Amagasu, S.M., Monroe, F.A. and MacIver, M.B., Halothane depresses action potential conduction in hippocampal axons, Brain Res. 796, 231–238, 1998. Xu, F., Sarti, P., Zhang, J. and Blanck, T.J.J., Halothane and isoflurane alter calcium dynamics in rat cerebrocortical synaptosomes, Anesth. Analg. 87, 701–710, 1998. Antkowiak, B., Different actions of general anesthetics on the firing patterns of neocortical neurons mediated by the GABAA receptor, Anesthesiology 91, 500–511, 1999. Nishikawa, K. and MacIver, M.B., Agent-selective effects of volatile anesthetics on GABAA receptor-mediated synaptic inhibition in hippocampal interneurons, Anesthesiology 94, 340–347, 2001. Li, X., Czajkowski, C. and Pearce, R.A., Rapid and direct modulation of GABAA receptors by halothane, Anesthesiology 92, 1366–1375, 2000. Ming, Z., Knapp, D.J., Mueller, R.A., Breese, G.R. and Criswell, H.E., Differential modulation of GABA- and NMDA-gated currents by ethanol and isoflurane in cultured rat cerebral cortical neurons, Brain Res. 920, 117–124, 2001. Kira, T., Harata, N., Sakata, T. and Akaike, N., Kinetics of sevoflurane action on GABA- and glycine-induced currents in acutely dissociated rat hippocampal neurons, Neuroscience 85, 383–394, 1998. Flood, P. and Coates, K.M., Sensitivity of the alpha7 nicotinic acetylcholine receptor to isoflurane may depend on receptor inactivation, Anesth. Analg. 95, 83–87, 2002. Flood, P. and Krasowski, M.D., Intravenous anesthetics differentially modulate ligand-gated ion channels, Anesthesiology 92, 1418–1425, 2000. Yamakura, T., Sakimura, K., Shimoji, K. and Mishina, M., Effects of propofol on various AMPA-, kainate- and NMDA-selective glutamate receptor channels expressed in Xenopus oocytes, Neurosci. Lett. 188, 187–190, 1995.
16 The Influence of Inhalational Anesthetics on Carotid Body Mediated Ventilatory Responses
ALBERT DAHAN, RAYMONDA ROMBERG, ELISE SARTON, and LUC J. TEPPEMA Leiden University Medical Center Leiden, The Netherlands
I.
Introduction
A major defense mechanism of the mammalian body to acute hypoxia is a rapid increase in inspiratory drive, the so-called acute hypoxic response (AHR). This vital and sometimes life-saving response is aimed at the enhancement of oxygen influx and originates at the peripheral chemoreceptors located in the carotid bodies (CB) [1–3]. In the sense that the CBs are the principal guards of adequate oxygen (and glucose; see Chapter 1 by Nurse, this volume) delivery to the brain, the CBs are strategically located at the bifurcation of the common carotid arteries [1]. The glomus type I cells of the CBs are thought to contain oxygen-sensing mechanisms. The full mechanism of oxygen sensing at the CBs is still poorly understood [2]. At present it is thought that membrane ion channels are critically involved and that low oxygen inhibits Kþ current through the CB type I cell membrane, which causes membrane depolarization and consequently the influx of calcium ions into the cell and the activation of a complex cascade of events within the type I cell ([2] and Chapter 1 by Nurse, this volume). At the end of this cascade, the cell releases neurotransmitters which activate 653
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postsynaptic receptors located on afferent fibers of the sinus nerve that have their cell bodies in the petrosal ganglion (PG) with their axons terminating in the nucleus tractus solitarii (NTS). In the PG, oxygen-sensitive autonomic and sensory neurons give rise to an extensive network of efferent fibers that innervate the CBs [4,5]. These neurons provide nitric oxide-mediated inhibition of the CBs when activated by hypoxia [5] and may play an important role in a negative feedback modulation of the hypoxic ventilatory drive. The peripheral chemoreceptors respond to hypoxia as well as to carbon dioxide and acute metabolic acidosis and, together with the central chemoreceptors located in the ventral medulla, play an important role in maintaining blood gas homeostasis (chemical or metabolic control of breathing) [6]. In the 1970s and early 1980s, the late Richard Knill and colleagues (Figure 16.1) at the University of Western Ontario in London, Ontario, were among the first to study the effects of low and high concentrations of the inhalational anesthetics on the control of breathing in humans [7–11]. Their studies on the older anesthetic agents halothane, enflurane, and isoflurane led, despite recent criticism, to the still generally accepted conclusion that inhalational anesthetics selectively impair all responses mediated by the peripheral chemoreceptors even at subanesthetic concentrations [MAC values of 0.1–0.2 MAC (minimum alveolar concentration or the anesthetic
Figure 16.1 The late Richard L. Knill (middle), Adrian W. Gelb (left) and research nurse Jane L. Clement, from the University of Western Ontaria at London, Ontario, Canada, perform an experiment on the influence of halothane on the ventilatory response to hypoxia (photo obtained with permission from Dr. Gelb).
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Table 16.1 Minimum Alveolar Concentration of Contemporary Volatile Anesthetics. Values are for 25- to 60-year-old Humans Enflurane Isoflurane Sevoflurane Desflurane Nitrous oxide
1.7% 1.25% 1.8% 7.0% 105%
concentration at which 50% of the study population does not respond (i.e., move) to a noxious stimulus such as skin incision or tail clamp)]. Human MAC values are given in Table 16.1. Note that MAC values 41 are needed for adequate anesthesia. Study of the influence of low- and high-dose anesthetics on CB-mediated responses in animals and humans is important, not only for possible clinical reasons, but because it may also help elucidate the mechanisms involved in oxygen sensing, locate the site of anesthetic action within the CBs (i.e., anesthetic-peripheral chemoreflex loop interaction as a model for anesthetic effect), and possibly lead to effective measures to prevent depression of CB-mediated responses by inhalational anesthetics. II.
Influence of Inhalational Anesthetics on the Ventilatory Response to Hypoxia and Hypercapnia
A. Ventilatory Response to Acute and Sustained Hypoxia
In adult humans, the ventilatory response to 15–20 min of hypoxia is biphasic (Figure 16.2) [12–18]. Under isocapnic conditions (i.e., end-tidal CO2 concentrations maintained constant), a stepwise reduction in end-tidal PO2 (PETO2 ) to mild hypoxia (45 mm Hg) that is sustained for more than 3– 5 min results in a rapid increase in ventilation followed by a slow decline (hypoxic ventilatory decline or HVD). A new steady-state ventilation, 25–40% above normoxic baseline, is observed within 15–40 min (Figure 16.2). Interestingly, HVD only develops when hypoxia is continuous. Multiple short-term hypoxic episodes (3-min hypoxia with 2-min normoxic interludes) are unable to generate HVD even when the cumulative duration of hypoxia exceeds 15 min [18]. In humans, the initial hyperventilatory response (acute hypoxic response or AHR) originates exclusively at the peripheral chemoreceptors in the CBs. Otherwise healthy patients show no ventilatory response to acute hypoxia or peripheral CO2 response after bilateral resection of their CBs ([19], see also below). The magnitude of the AHR is variable among normal subjects and patients, ranging from 0.1 to 2.0 l/min per % hemoglobin– oxygen desaturation [20]. When isocapnia is not maintained (i.e.,
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Hypoxia
A 35
Ventilation (l/min)
30 25 20 15 10 5 0 B 35 Ventilation (l/min)
30 25 20 15 10 5 0 0
5
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40
Dopamine Time (min)
Figure 16.2 Ventilatory response to sustained isocapnic hypoxia followed by a short normoxic episode and a second exposure to isocapnic hypoxia. A. The hyperventilatory response is followed by a slow ventilatory decline due to the depressant effect of central hypoxia. Due to the ongoing hypoxic depression, a next hypoxic response is suppressed by 50%. B. Low-dose dopamine (3 mg kg1 min1) suppresses the hypoxic drive at the CBs. Despite central hypoxia, a second hypoxic response, now without any dopamine, is not depressed. This indicates the need for afferent input from the CBs during sustained hypoxia for hypoxic ventilatory depression to develop. Each data point is a 2-min average. Data are from one subject (# 96_022) (from Ref. 18).
poikilocapnia), the ventilatory response is much smaller (0–0.5 l/min per % desaturation). Both human and animal studies show that upon sudden exposure to hypoxia, the hyperventilatory response has a fast component (time constant 2–6 sec) and a slow component (time constant about 60–120 sec) [21–25]. It is thought that the fast component is the true CB response, while the slow component is due to the central modulation of the CB response
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(i.e., central neuronal dynamics or short-term potentiation of breathing, STP) [21]. Note that activation of the CBs with carbon dioxide does not produce a similar (two-component) pattern in ventilation [26]. The mechanism(s) and site(s) of generation of HVD in awake humans remain poorly understood. There are several possibilities: 1.
Adaptation of the CB activity during sustained hypoxia (i.e., an exclusive peripheral effect) [27]; 2. Mechanisms within the central nervous system (CNS) related to hypoxia-induced changes in brain blood flow (BBF) and/or the build-up of substances with a net inhibitory effect on breathing during central hypoxia (i.e., an exclusive central effect) [28–31]; 3. The modulation within the CNS of the peripheral drive from the CBs into HVD (i.e., peripheral-central interaction) [17,29]. Studies in anesthetized cats suggest that adaptation of the peripheral drive from the CBs during sustained hypoxia is highly unlikely to be a major mechanism of HVD. Vizek et al. [3] and Andronikou et al. [32] showed, in anesthetized cats, a fast increase in carotid sinus nerve activity upon hypoxic exposure. In contrast to phrenic nerve activity, sinus nerve activity remained unaltered for at least 20 min. Indications that the CNS is exclusively responsible for the development of HVD come from a study in cats using the technique of artificial brainstem perfusion [28]. When hypoxia was limited to the brainstem, a decline in ventilation was observed. Furthermore, the dynamics of these ventilatory changes were similar to those observed after changes in end-tidal CO2 (time constant 2 min). This suggests, at least in the anesthetized cat, that a hypoxiainduced increase in BBF causing the washout of acid metabolites (CO2, Hþ) from the brain is the main mechanism of HVD. In awake humans, part of HVD is related to the increase in BBF during hypoxia. In awake humans, there is a decrease in the gradient between jugular venous and arterial PCO2 of 2 mm Hg [14], which, taking into account the slope of the hypercapnic ventilatory response (1–2 l/min per mm Hg [14]), is sufficient for 15–30% of HVD [15]. A more important mechanism in awake humans may be the central accumulation of inhibitory neuromodulators/transmitters such as adenosine, g-amino butyric acid (GABA) or dopamine, causing a net inhibitory effect on the respiratory neuronal pool in the brainstem [29–31]. The relatively slow turnover of these substances after the relief of hypoxia may explain the persistent reduction of hypoxic ventilatory responses after 20-min hypoxic exposures (see also Figure 16.2, top panel), which needs at least one hour to wane [13,17]. Interestingly, in awake humans and animals, central hypoxia in the absence of a hypoxic drive from the CBs (for example, in bilateral CB-resected subjects or due to silencing of the CBs with low-dose dopamine) is unable to generate HVD or depress subsequent hypoxic responses (Figure 16.2)
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[17,33–35]. This suggests in awake mammals that the increased afferent input from the CBs during hypoxia activates the buildup of inhibitory agents in the CNS (possibility 3 above). Animal studies show that CB stimulation caused by hypoxia or hypercapnia in turn causes the increase of glutamate (a stimulatory neurotransmitter) in respiratory centers in the CNS [36,37]. Glutamate serves as the precursor of GABA; conversion of glutamate into GABA during sustained hypoxia may cause a net inhibitory effect on ventilation (i.e., HVD) [38]. This mechanism not only explains the lack of HVD in CB-resected subjects, but also the observation that the magnitude of HVD is proportional to the magnitude of the acute ventilatory response to hypoxia (increasing/decreasing AHR using specific agents such as almitrine/dopamine or CB resection causes greater/lesser HVD) [17,33–35,39]. The picture that emerges from the literature is that in awake adult mammals the development of HVD requires central (i.e., within the CNS) and peripheral hypoxia combined with an intact hypoxic drive from the CBs. The hypoxic drive plays a crucial facilitatory role in the development of HVD. In this sense, HVD is a CB-mediated response. In anesthetized humans and animals, HVD is mainly related to hypoxic-induced increases in brain blood flow. B. Human Studies Halogenated Volatile Anesthetics at Subanesthetic Concentrations
The halogenated anesthetic agents are an example of a set of drugs that, at anesthetic concentrations, affect ventilatory control at a number of sites. For example, halothane at anesthetic concentrations (1 MAC) causes ventilatory depression by abolishing the peripheral drive of the CBs (the ventilatory response to two breaths of O2 is absent at 1 MAC [40]), by changing the balance between excitatory and inhibitory neuromodulators within the central nervous system toward a net inhibitory effect on the respiratory neuronal pool (see Chapter 15 by Stuth, this volume), and by suppression of respiratory muscles [diaphragm and intercostal muscles ([41] and see Chapter 17 by Warner, this volume)]. Furthermore and clinically important, this loss of CB function will lead to a decrease in ventilation (i.e., HVD) without any hyperventilatory response upon the exposure to hypoxia at halothane concentrations of about 1.1 MAC [7]. The ventilatory response to acute hypoxia
Knill’s studies on hypoxic and hypercapnic ventilatory responses indicated a powerful depressant effect of halothane, enflurane, and isoflurane at just 0.1 MAC on the ventilatory response to a gradual decrease in oxygen saturation, while little to no effect was observed on the ventilatory response to CO2 as assessed by Read’s rebreathing technique [7–11]. The results of
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Knill’s studies lead to the still generally held conclusion that, . . . in humans, all halogenated anesthetic agents at subanesthetic concentrations cause the effective loss of peripheral chemoreflex-mediated functions in the presence of hypoxemia, (which causes) not only the loss of physiological defenses but also of clinical signs [42]. This statement was made in 1982 when arterial oxygen saturation could not be monitored continuously in an economical and reliable way, something clinicians today cannot image. However, today this statement retains its importance for large groups of perioperative patients. It took several years before five independent groups from Europe and the USA undertook the replication of Knill’s studies [16,20,23,24,45–58]. At present, there are more than 25 separate studies on the influence of low-dose anesthetics on ventilatory control in humans. Although some differences in results are observed (Table 16.2), the overall picture that emerges from the literature is that the halogenated agents halothane, enflurane, isoflurane, and sevoflurane do affect the hypoxic ventilatory response with distinct potencies [59]. Differences in study results among the various groups may be related to differences in study protocol such as the presence/absence of isocapnia within and among studies and differences in the behavioral state of the subjects while inhaling the anesthetic agents (see Table 16.2 and Section III, below). Differences in the methods of inducing hypoxia could not explain any of the observed differences in study outcomes. Both step hypoxic tests, in which hypoxia is rapidly induced (i.e., within 4–6 breaths) and maintained constant for at least 3 min, and ramp hypoxic tests, in which hypoxia gradually develops over 10–15 min, give similar results (Figure 16.3) [44]. This is surprising considering that HVD develops during the 10–15 min of hypoxia and inhalational anesthetics may enhance HVD (Figure 16.4A and Figure 16.4B). Halothane is the most potent depressant of the AHR (450% depression at 0.1 MAC), followed by enflurane (50% depression), sevoflurane (30% depression at 0.1 MAC) and isoflurane (20–40% depression at 0.1 MAC) (Table 16.2) [59]. Lesser depression may be observed when the subjects are aroused and behavioral control is activated (see Section III below). Desflurane seems to be an exception to the rule, causing no depression of the AHR at 0.1 MAC at normocapnia (PETCO2 43 mm Hg), although 30% depression is observed when hypoxia is combined with hypercapnia (PETCO2 50 mm Hg) [20]. The differences in potency to depress AHR among these anesthetics may be related to activation of the sympathetic system by some anesthetics (e.g., isoflurane and desflurane), thus counteracting part of the depression of the AHR, or may be more fundamental and involve differences in the interaction of the anesthetics with the complex process of oxygen sensing at the CBs (see below). Note finally that halothane’s and isoflurane’s dose-AHR relationships are
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Table 16.2 An Overview of Studies from the Literature on the Influence of Subanesthetic Halothane, Isoflurane, Desflurane and Sevoflurane on the Acute Ventilatory Response to Hypoxia (AHR) Institution Halothane University of Western Ontario LUMCa
Oxford University
University of Edinburgh Isoflurane University of Western Ontario UCLAb LUMC
Karolinska Institute University of Edinburgh Desflurane LUMC Sevoflurane LUMC
University of Oxford a
Concentration
n AHR Isocapnia Ref. (year)
0.05 MAC 0.1 MAC 0.08% inspired 0.15% inspired 0.22% inspired 0.11% end-tidal 0.05 MAC 0.1 MAC 0.2 MAC 0.1% inspired
10 10 9 9 10 10 12 12 12 9
0.50 0.30 0.70 0.32 0.32 0.45 0.65 0.50 0.25 0.70
Yes Yes Yes Yes Yes Yes Yes Yes Yes No
[7] (1978) [7] (1978) [45] (1994) [45] (1994) [16] (1994) [57] (2002) [47] (1995) [47] (1995) [47] (1995) [46] (1993)
0.1 MAC 0.1 MAC 0.1 MAC 0.1 MAC 0.1 MAC 0.2 MAC 0.11% end-tidal 0.2% end-tidal 0.1 MAC 0.1 MAC
12 8 8 10 8 8 10 18 20 9
0.50 0.94 0.85 0.50 0.50 0.36 0.60 1.00 0.70 1.20
Yes Yes Yes Yes Yes Yes Yes No Yes No
[9] [43] [44] [48] [49] [49] [58] [50] [51] [52]
14 0.94
Yes
[20] (1996)
11 20 9 9 8
Yes Yes Yes Yes Yes
[53] [54] [55] [55] [56]
0.1 MAC 0.1 MAC 0.25% end-tidal 0.14% end-tidal 0.27% end-tidal 0.1 MAC
0.70 0.64 0.75 0.50 0.80
(1983) (1992) (1994) (1994) (1995) (1995) (2002) (1994) (1995) (1995)
(1996) (1999) (2000) (2000) (1999)
Leiden University Medical Center. University of California Los Angeles.
b
particularly steep and small errors in end-tidal anesthetic concentration measurements may be the cause of large errors in the presumed depression of the AHR (Figure 16.4C and Figure 16.4D). The utilization of a calibrated vaporizer and anesthetic measurement device, and the use of precise gas flows (through the vaporizer and delivered to the subject), may significantly reduce but not completely abolish this error. Evidently, this may be another cause of the differences in study outcomes observed by the different research groups (Table 16.2).
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A. Step hypoxic tests
30
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. Vl (l/min)
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0 0
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O2
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[100 − SpO2] (%)
Figure 16.3 Influence of 0.2% end-tidal isoflurane on the ventilatory response to isocapnic hypoxia assessed by two different methods. A. Steady-state response obtained at five different end-tidal PO2 levels using step hypoxic inputs (duration of each hypoxic step was 3 min). The lines through the data are fits to the equation VI ¼ A exp(D PETO2 ). B. Nonsteady-state response obtained by using a pseudorebreathing technique in which hypoxia gradually developed over 10–12 min. The lines through the data are linear regression fits. The response slope (hypoxic sensitivities) is given. A and B. Open symbols are control data, closed symbols isoflurane data. In both studies the effect of isoflurane was of similar magnitude, causing a 60% depression of the ventilatory response to hypoxia. SpO2 is arterial oxygen–hemoglobin saturation derived from pulse oximetry. Data are from one subject (# 98_002) (Unpublished data from Dahan, 1997).
While only animal studies can give definite proof, there is now ample albeit indirect evidence that the site of action of low-dose inhalational anesthetics in humans is at the peripheral chemoreceptors of the CBs: Hypoxia-driven ventilation decreases by 25% within 30 sec and by 40% within 1 min of exposure to 0.15–0.30% inspired halothane. Within this time period, brain concentrations of halothane cannot contribute to this effect, given the relatively slow blood-effect-site equilibration of halothane (at 30 sec of halothane-wash-in, brain concentrations will be 2% or less of end-tidal, while CB tension will be 70% of end-tidal; at 60 sec of wash-in these values are 10% and 90%) [7]. Hypoxia-driven ventilation (arterial O2–hemoglobin saturation 80%) decreases by about 3 l/min within 30 sec upon exposure to isoflurane (end-tidal isoflurane concentration ¼ 1 mm Hg). Taking into account the time constant for isoflurane uptake in CBs and the brain (7 sec and 4 min, respectively), the CB isoflurane tension is 90% of end-tidal, whereas the brain concentration is just 8% of end-tidal [48].
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A
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AHR (% of baseline ventilation)
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0 2 4 6 8 10 12 14 16 AHR (l/min)
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200 180 160 140 120 100
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0.0 0.2 0.6 0.4 0.8 1.0 1.2
0.0 0.2 0.4 0.6 0.8 1.0 1.2
Fraction of MAC halothane
Fraction of MAC isoflurane
Figure 16.4 A. Influence of 0.15 MAC halothane on the ventilatory response to sustained isocapnic hypoxia. Despite the lesser initial hyperventilatory response (from 22 to 9 l/min), the development of hypoxic ventilatory decline remains unaffected (from 10 to 9 l/min). Note the sharp undershoot after the relief of hypoxia by normoxia during halothane inhalation (6 l/min) but not during the control study (1 l/min). Each data point is a 2-min average. Data are from one subject (# 94_077) (from Ref. 48). B. Influence of 0.15 MAC halothane on the relationship between the AHR and hypoxic ventilatory depression (HVD). Note the facilitation of HVD development by halothane (from Ref. 48). C and D. Dose-response for the effects of halothane and isoflurane on the AHR (data from Refs. 45, 48 and 49).
Halothane at 0.1 MAC depresses by 60% the ventilatory response to acute metabolic acidosis via infusion of l-arginine hydrochloride (acute, moderate metabolic acidosis affects ventilation mainly through stimulation of the CBs) [11]. Using the dynamic-end tidal forcing technique, the ventilatory response to a square-wave change in end-tidal PCO2 may be
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separated into a fast component arising from the peripheral chemoreceptors and a slow component arising from the central chemoreceptors. This noninvasive technique gives the most crucial evidence for a preferential, peripheral effect of low-dose inhalational anesthetics. Halothane, isoflurane, and sevoflurane at 0.1 MAC reduced the peripheral CO2 sensitivity without affecting central CO2 sensitivity [45,49,60]. The ventilatory response to sustained hypoxia
There is only a handful of studies on the influence of inhalational anesthetics on the development of HVD [16,43,46,49,52,55,61]. Knill showed that at anesthetic concentrations, halothane produces HVD despite the absence of AHR [7]. This is surprising taking into account the observation that AHR is required for development of HVD in awake humans, and indicates a distinct mechanism of HVD development during anesthesia. Berkenbosch and colleagues, using the artificial brainstem perfusion technique, showed the development of HVD during exposure to hypoxia limited to the CNS in a-chloralose/urethane anesthetized cats [28]. Both observations indicate that under conditions of anesthesia, the coupling of AHR and HVD is lost in cats and humans. The observation that the dynamics of the slow ventilatory roll-off during sustained hypoxia is similar to that of central CO2 sensitivity (both about 2 min) suggests an important role for BBF in the development of HVD under anesthetic conditions [28]. In addition, during the inhalation of 0.15 MAC halothane (but not isoflurane, enflurane and sevoflurane), the awake coupling between AHR and HVD is changed with a leftward-shift of the relationship indicating the need for less AHR to develop HVD (Figure 16.4B) [16]. Since at these inhaled concentrations (50.6 MAC) halothane does not affect BBF, it is unlikely that the relative increase in HVD is related to halothane-hypoxic interaction on BBF. A more plausible mechanism is the synergistic increase of inhibitory neuromodulators during hypoxia/halothane exposure [62].
Molecular Mechanism of Anesthetic Effect and Involvement of Radical Oxygen Species (ROS) in Anesthesia-induced Depression of the Hypoxic Response
How anesthetics produce their state of unconsciousness is unknown, but several likely molecular targets have been identified. The most promising candidates are inhibitory ligand-gated ion-channels, such as GABAA and glycine receptors [63]. Recent studies indicate that certain mammalian (including human) background Kþ channels are activated by general anesthetics and are thought to be involved in the side effects of anesthesia such as cardiac depression (bradycardia and negative inotropic effects) as
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well as respiratory depression [64]. These Kþ channels belong to a gene family in which each subunit has a unique structural motif of two poreforming domains in tandem plus four transmembrane segments (2P/4TM) and, at present, consists of a handful of members (TREK-1, TRAAK, TWIK-1, TASK, and TALK-channels) [63]. For the TASK channel, the letters indicate tandem P domain acid sensitive Kþ. The mechanisms of oxygen sensing at the CB and the mechanism(s) through which inhalational anesthetics disrupt O2 sensing remain poorly understood, but most probably involve a type I cell membrane ion and/or cytoplasmatic (non)-mitochondrial heme-containing enzymes [1,2,65]. Results from in vivo and in vitro studies indicate that potassium channels (e.g., TASK, Kv, maxi-K channels) initiate the oxygen-sensing transduction cascade [1,2,65–70]. The exact mechanism through which low oxygen increases the conductance of Kþ channels is unknown, but may be related to their sensitivity to reactive oxygen species (ROS) and to changes in redox state [71,72]. Reactive species themselves, on the other hand, may also be involved in oxygen sensing but presently their exact role remains obscure [72]. It is known that in hypoxic conditions, volatile anesthetics (particularly halothane) produce reactive species. This action causes lipid peroxidation and mild liver damage, an effect that in guinea pigs can be prevented by antioxidant treatment [73–77]. Also, volatile anesthetics such as halothane increase the conductance of potassium, especially TASK-channels, possibly by binding to a specific cytoplasmatic c-terminal domain that is also required for neurotransmitter inhibition of the channel [63,64,78–80]. The findings that low oxygen closes potassium channels which by themselves are sensitive to ROS, and that halothane is not only able to open potassium channels but also produces ROS (particularly in hypoxia), raise the question whether halothane may reduce the hypoxic response by producing ROS and/or by influencing the redox state of the CB type I cell. We investigated this issue in healthy volunteers by studying the influence of an antioxidant cocktail (a-tocopherol and ascorbic acid) on the AHR [57]. The antioxidants prevented depression of the hypoxic response by 0.13 MAC halothane (Figure 16.5A). In fact, none of the subjects pretreated with antioxidants showed any depression of the ventilatory response by halothane, while subjects pretreated with placebo showed 50–60% depression. Antioxidants by themselves had no effect on AHR. The finding that antioxidants effectively prevented the depression of the AHR by subanesthetic halothane indicates that this agent acts through reactive species generated by its reductive metabolism. An increase in ROS concentration would then depress the hypoxic response by opening ROSsensitive potassium channels. An alternative explanation could be that the antioxidants, by changing the cellular redox state or the concentration of
B. Isoflurane 100
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AOX 90
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CON
ISO
ISO + AOX/PL
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Figure 16.5 The ability of antioxidant pretreatment (a combination of oral a-tocopherol and IV ascorbic acid) to prevent halothane(A. left) and isoflurane- (B. right) induced depression of the ventilatory response to acute isocapnic hypoxia in healthy volunteers. AOX is the antioxidant group, PL is the placebo group. In the halothane study, AOX and PL were two separate groups of eight subjects; the isoflurane data is paired (n ¼ 10). A. Halothane and halothane þ placebo: P 5 0.01 versus control and halothane þ antioxidants. B. Isoflurane and isoflurane þ placebo P 5 0.01 versus control and isoflurane þ antioxidants (Data from Refs. 47 and 48).
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ROS, may prevent or modify the binding of halothane to the specific domain of the potassium channel. Alternative sites of action of the volatiles also need to be considered, such as complexes I or II of the respiratory chain in mitochondrial membranes, that are molecular targets of volatile anesthetics (Figure 16.6) [81] and are possibly involved in the O2 sensing cascade [82,83]. Some insight into the involvement of complexes I and II in O2 sensing may be obtained from data collected in patients with specific mutations in the different genes that encode the cytochrome proteins. For example, the mitochondrial respiratory chain complex II (which consists of four subunits: succinate dehydrogenase (also known as succinate ubiquinone oxireductase) A, B, C and D or SDHA, SDHB, SDHC and SDHD) is involved in the Krebs cycle (by catalyzing the oxidation of succinate to fumarate) and in the mitochondrial electron transport chain [83–85]. Complex II is anchored to the mitochondrial membrane by the polypeptides SDHC and SDHD. SDHA is a flavoprotein (the active site of the enzyme), while SDHB is an Fe-S subunit, both projecting into the mitochondrial matrix [81]. A missense mutation in the gene that encodes for SDHD (PGL-1 mutation in the SDHD gene on chromosome 11q23, leading to the change of aminoacid residue ASP92 to Tyr in the gene product) leads to slow-growing paraganglionic tumors (including tumors of the CBs and pheochromocytomas) in Dutch founder families [84,84a]. The PGL-1 mutation may lead to reduction in SDH enzyme activity, reduced energy (ATP) production, and oxygen free radical formation [85]. We recently measured the hypoxic ventilatory sensitivity in heterozygous PGL-1 positive subjects from these families who were still without Halothane Isoflurane Sevoflurane NADH:Q oxidoreductase
N2O Cyt. c
Q
Complex I
Complex III
O2 Complex IV
Halothane Succinate dehydrogenase Complex II
Figure 16.6 Sites of action of inhalational anesthetics within the mitochondrial respiratory chain. Halothane affects both complex I and II, while sevoflurane and isoflurane affect complex I selectively (Data from Ref. 81).
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CB tumor development. The hypoxic sensitivity tended to be 40–60% smaller than responses obtained in controls, but was still within the normal range observed in the general population. Most importantly, these same subjects have normal peripheral and central CO2 sensitivities but lack the characteristic peripheral O2–CO2 interaction. In other words, subjects with the PGL-1 gene mutation showed no multiplicative increase in peripheral CO2 sensitivity from normoxia to mild hypoxia. Our findings suggest a (modulatory) role for complex II in O2 sensing and O2–CO2 interaction. Further study is needed to determine whether this is related to excessive or aberrant radical oxygen species formation (related to the disruption of electron transport in the cytochromes) and/or dismutation, and whether complexes I and II are the sites of action for anesthesia-induced depression of the AHR (inhalational anesthetics may then cause disruption of the electron transport within the mitochondrial respiratory chain, causing the accumulation of ROS in the cell cytoplasm with subsequent modulation by ROS of the primary oxygen sensing site). Finally, we further observed in healthy volunteers that antioxidants were also able to prevent the 40% depression of the AHR by 0.1 MAC isoflurane, a volatile anesthetic that is also known to produce reactive species, albeit to a lesser degree than halothane (note that the depression from isoflurane is also smaller than from halothane; Figure 16.4D, Figure 16.5B and Figure 16.7) [58]. Desflurane (0.1 MAC), on the other hand, has a very low metabolism with very little production of reactive species and does not impair the hypoxic response (see above and Figure 16.7) [20]. It is also interesting to note that the intravenous anesthetic propofol, which has antioxidant properties, depressed neither the CO2 sensitivity of the peripheral chemoreflex loop nor the fast, CB-mediated component of the hypoxic response (see below) [86]. Together, these findings in animals and humans suggest that the depressant effect of anesthetics on the hypoxic response may be related to their pro-oxidant properties, but further studies are needed to confirm or exclude this. C. Animal Studies
In most animal preparations, the inhalational anesthetics inhibit the ventilatory responses to hypoxia and hypercapnia [87–97]. However, animal studies are equivocal with respect to sites of anesthetic action. Although all investigators agree that the ventilatory control system is depressed at pathways common to both chemoreflex loops, (1) some have found an additional selective effect within the peripheral chemoreflex loop at the CBs ([87–91], see below); (2) some have not ([92–95], see below); and
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Isoflurane
Sevoflurane
60 Enflurane 50 Halothane
40 30 0.01
0.1
1
10
100
Metabolism (%)
Figure 16.7 Relative metabolism of the inhalational anesthetics in humans versus their depressant effect on the acute hypoxic ventilatory response at 0.1 MAC (AHR data from Ref. 59).
(3) others have shown a dependency of a selective effect on the hypoxic stimulus intensity ([96], see below). Regarding item 1, selective effect at the CBs: In dogs, Weiskopf et al. showed that 1.1% halothane impaired the ventilatory response to hypoxia to a greater extent than the response to CO2 and interfered with the synergistic interaction of hypoxia and hypercapnia on ventilation [87]. In a different species, the cat, Davies et al. measured the effect of 0.5– 1.0% halothane on single- or few-fiber preparations of the carotid sinus nerve in decerebrated animals. The slopes of the CB response to both hypoxia and hypercapnia were depressed. The halothane response was prompt (half-life 1–2 min) and reversible [88]. In vagotomized dogs, Stuth et al. measured the effects of halothane on the phrenic nerve response to 1–1.5 sec infusions of 100% CO2 in saline into the carotid arteries. Halothane caused a dose-dependent (0.5–2 MAC) reduction in phrenic nerve activity, which was not seen after bilateral CB resection [89]. In anesthetized (pentobarbital and a-chloralose) and artificially ventilated cats, Ide et al. studied the influence of halothane on phrenic
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nerve responses to hypoxia. The CBs were selectively perfused with a hypoxic solution containing 0, 0.1 or 0.8% halothane. The authors showed a dose-dependent depression of the hypoxic response and complete abolition of the response at 0.8% [90]. In an anesthetized (a-chloralose–urethane) cat model, using squarewave changes in end-tidal CO2, sevoflurane caused a dosedependent depression of the peripheral chemoreflex loop of greater magnitude than the depression of the central chemoreflex loop in a-chloralose and urethane anesthetized cats [91]. Regarding item 2, absence of selective effect at the CBs: Using the artificial brainstem perfusion technique, the effect on the ventilatory responses to hypoxia and hypercapnia of central (brainstem) versus peripheral and overall (central þ peripheral) halothane was studied in a-chloralose and urethane anesthetized cats. Central and peripheral CO2 sensitivities were depressed by 0.5–1.5% halothane without changing the ratio between these sensitivities. The ventilatory response to hypoxia was depressed independently of the perfusion condition. The authors concluded that although a direct effect of halothane on the peripheral chemoreceptors cannot be excluded, such an effect might be of minor importance in comparison with the depressant effect of halothane on other peripheral structures, i.e., the neuromechanical link between brainstem and ventilation [92,93]. In yet another species, the awake goat, Koh and Severinghaus measured the ventilatory responses to CO2 and hypoxia at 0.5– 1.25% halothane. At 0.5% end-tidal concentration, halothane did not affect hypoxic chemosensitivity. At higher concentrations CO2 and hypoxic responses were affected similarly [94]. In vagotomized dogs, Stuth et al. tested the influence of halothane on the phrenic nerve response to hyperoxia and hypoxia. Halothane caused a dose-dependent depression of hypoxic responses, but did not abolish the response at surgical concentrations (2 MAC). Relative hyperoxic and hypoxic phrenic nerve activities were depressed similarly at the different halothane concentrations [i.e., a parallel shift of phrenic nerve activity at the different halothane concentrations (0.5–1 MAC) for the two O2 states] [95]. Regarding item 3, Dependency on stimulus strength. Finally in the rabbit, Ponte and Sadler measured carotid sinus nerve neural activity during halothane, isoflurane, and enflurane inhalation and different arterial oxygen concentrations. In doses up to 1%, the steady-state chemoreceptor discharge response to hypoxia
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Human
+++
Cat Depression of AHR @ 0.5 % halothane Rabbit
Goat
0 0
+++ Production of ascorbic acid
Figure 16.8 Relationship between the production of ascorbic acid and the depression of the ventilatory response to acute isocapnic hypoxia by 0.5% end-tidal halothane in four species. Humans have lost their ability to produce the antioxidant and have the greatest depression of the hypoxic response by halothane.
was shifted downward. However, hypoxic stimuli below 40 mm Hg overcame this depressant effect [96]. Evidently, the variability in study outcomes may be related to species differences; methodological differences with respect to anesthesia (e.g., the presence or absence of background anesthesia or decerebration); differences in surgical preparations (e.g., artificial brain stem perfusion versus phrenic nerve reading in vagotomized animals); absence of dose-response curves in some studies; possible masking of peripheral responses by systemic hypotension, and differences in hypoxic stimulus intensities. Finally, it is of interest to speculate whether the variability in halothane-induced depression of the AHR among different species may be related to ROS. While in goats an end-tidal concentration of 0.5% halothane does not significantly depress the hypoxic drive [94], similar concentrations do cause depression in cats and rabbits, with a greater effect in cats [84,96]. These differences could be related to differences in the oxygen-sensing mechanisms (for example, the type of oxygen-sensitive potassium channel, or the expression of a variety of channel splice variants among the species with differences in anesthetic sensitivities). An alternative explanation could be related to species differences in defense mechanisms against ROS (Figure 16.8). Note
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that goats produce large quantities of ascorbic acid and may thus be better protected against the adverse effects of ROS and free radicals and consequently also against the adverse effects of halothane [98]. To a lesser degree this is also true for rabbits. Cats produce low quantities of ascorbic acid and this might explain their higher susceptibility to halothane than that of rabbits [98]. Humans have lost the ability to synthesize ascorbic acid and may therefore be more vulnerable to the adverse effects of ROS (Figure 16.8). [Note that anesthetic potency is somewhat lower in animals when compared with humans. For example, canine and feline MAC values are 2.4% for sevoflurane and 0.9% for halothane (compare with human values in Table 16.1). In conclusion, we postulate a hypothesis that explains the variability in effect of the various inhalational anesthetics on the CB response to hypoxia in a single species (the human) as well as among different species, and explains the absence of CB effect of propofol, which has antioxidant properties. This unified theory implicates a species- and anesthetic-independent key role for ROS in anesthesia-induced depression of the hypoxic drive. III.
Pain and Behavioral Responses
Inspired minute ventilation is under the influence of various factors and two important control systems [99]. Most important is the chemical or metabolic control of breathing (that is, control of breathing solely dependent on the chemical composition of the arterial blood), which operates exclusively during light non-REM sleep and anesthesia. Behavioral control, which depends on nonchemoreceptor input, can modulate or even temporarily override the metabolic system. As a result, breathing is adapted to various circumstances such as talking, singing, playing a musical instrument, exercising, diving, etc. Behavioral control is also activated/ inhibited by various stimuli, such as pain, stress, arousal, sedation, and loss of consciousness, which occur in the perioperative phase. Pain interacts with ventilatory control via interaction with respiratory centers in the pons and medulla (pain-related control of ventilation) [53]; sedation/arousal causes control from higher centers via cortico-spinal pathways (acting directly at the respiratory motoneurons) or via the reticular system (suprapontine control of ventilation) [100]. Under normal study conditions (the subject is relaxed and not aroused), tests that assess the ventilatory response to inspired carbon dioxide are predominantly chemical in nature, while resting parameters (resting ventilation and end-tidal PCO2) have strong behavioral and chemical components. Clinical and experimental studies show that pain and surgical stimulation act as respiratory stimulants in the awake, sedated, and anesthetized states [53,101–104]. In terms of ventilation, pain is able to
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Figure 16.9 Influence of acute pain on the ventilatory response to isocapnic hypoxia in the awake state (A) and during 0.1 MAC sevoflurane inhalation (B) in one healthy volunteer. Note the pain-induced chemoreflex-independent ventilatory drive, which was independent of the arousal state of the subject. During sevoflurane inhalation and noxious stimulation, the subject was wide awake with open eyes. Each data point is the mean of 2 min (Data from Ref. 54).
reverse the respiratory depression from anesthetics and opioids. However, the effect of pain on respiration is chemoreflex-independent [53,104]. For example, experimental moderate pain (visual analog scale ¼ 5), induced by activation of cutaneous nociceptors on the lower extremities, did not affect 0.1 MAC sevoflurane-induced depression of the AHR (despite fully awake subjects), but did shift the response to higher ventilation levels due to an increase in pre-hypoxic baseline ventilation (Figure 16.9) [53]. Similar observations are made at higher anesthetic concentrations with more intensely painful stimulation (surgical pain) [101,102], indicating that pain is unable to reverse anesthetic-induced impairment of chemoreflex-related responses. From a clinical point of view, all that matters is whether a patient maintains an adequate minute ventilation. Since pain increases ventilatory drive it may be partly able to offset the anesthetic-induced loss of chemoreceptor drive.
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Figure 16.10 A. Influence of 0.1 MAC isoflurane on the ventilatory response to acute isocapnic hypoxia in a subject at rest without any tactile, audio or visual (AV) stimulation and with closed eyes. Note the depression of the response during isoflurane inhalation. B. The same subject now watching a music video and not allowed to close his eyes. The depression of the AHR is no longer present due to activation of behavioral control of breathing. Each data point is the mean of 1 min (Data from Ref. 48).
In contrast to pain, arousal responses due to vigorous tactile, audio and visual stimulation do interact with the peripheral chemoreflexes [43,48]. When humans are aroused by any of these stimuli, the chemoreflex response to hypoxia remains intact or is increased (Figure 16.10). The site at which arousal interacts with pathways involved in CB responses to hypoxia remains unknown but may be at central (see above) or peripheral sites, and may in both cases be related to the increase in catecholamine levels during vigorous stimulation. An interesting observation in this respect is that the neurotransmitters noradrenaline (NA) and serotonin (5HT) and the anesthetic halothane converge on TASK channels with opposite effects. Both NA and 5HT were able to reverse halothane-induced hyperpolarization and increased neuronal excitability [105]. It follows from these observations that specific experimental conditions are required when testing the influence of low- and high-dose anesthetics on the chemical control of breathing (quiet and comfortable surrounding, absence of tactile, audio and visual stimulation, empty bladder, experienced subjects, etc.). These observations
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explain most of the differences in the outcome of the studies of isoflurane by Temp, Knill and others (Table 16.1) [9,43,44]. IV.
Short-Term Potentiation of Breathing (STP)
There is ample evidence that chemoreceptor-mediated ventilatory responses undergo modulation within the CNS, resulting in respiratory activity not related to the dynamics of the chemoreceptors per se, and is called shortterm potentiation (STP) of breathing [106]. Synonyms include ventilatory afterdischarge, central neuronal dynamics, and respiratory memory. It has been suggested that STP plays an important role in stabilizing ventilation when ventilatory drive is suddenly reduced and prevents periodic breathing and apnea [107]. The occurrence of STP is well documented in animals. In cats, a rapid initial fall in inspiratory activity at the termination of electrical stimulation of the CB sinus nerve was observed, followed by a slow-decaying afterdischarge [108]. In the same species, a lag was observed between the changes in phrenic nerve activity and medullary extracellular fluid [Hþ] after a square-wave change in arterial or end-tidal PCO2 [106,109]. Finally in cats, the ventilatory response to a change in end-tidal PO2 with a background of constant arterial medullary PO2 is best described by a two-compartment model with time constants of 2 and 60 sec. The slow component is considered to be the manifestation of STP [21]. In humans, STP exhibits slow dynamics and is observed during stimulation of the (central and peripheral) chemoreceptors and suprapontine centers [22–25]. In awake humans, 60 sec of isocapnic hypoxia causes a ventilatory response, which is best described by two components with time constants ( ) of 4 sec and 30 sec [23–25,86]. An example of STP activation is given in Figure 16.11A. Upon relief of hypoxia, the manifestation of STP as a slow ventilatory decay is obscured but not abolished by a hypocapnic background, prior sustained hypoxia, and the inhalation of a hyperoxic gas mixture following hypoxia [25]. This statement is appreciated when considering the effect of hyperoxia on STP. Hyperoxia following 30–60 sec of hypoxia causes the loss of hypoxic drive followed immediately by a stimulatory effect on breathing due to the reduced Haldane effect and some decrease in BBF. As a consequence, after the fast drop in ventilation (i.e., the loss of hypoxic drive), the slow decrease in ventilation (STP) remains unobserved [25]. In humans, there are several indications that inhalational anesthetics inactivate STP: In volunteers, inhalation of 50 and 75% of the inhalational anesthetic nitrous oxide causes hypoventilation, apnea, and oxygen desaturation after 2 min of poikilocapnic voluntary hyperventilation [110,111].
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A. Awake control
B. 0.1 MAC isoflurane
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Figure 16.11 The ventilatory pattern following 60 sec of isocapnic hypoxia in one subject. A. In the awake control state, a fast and a slow ventilatory decline are observed with time constants of 3 and 40 sec, respectively. The slow decline is the manifestation of short-term potentiation of breathing. B. During the inhalation of 0.1 MAC isoflurane, only a fast ventilatory decline towards prehypoxic baseline ventilation is observed. The short-term potentiation of breathing is inactivated by the inhalational anesthetic (Data from Ref. 24).
Upon the termination of 1 min of isocapnic hypoxia, only one decaying ventilatory component was observed during 0.1 MAC halothane or isoflurane inhalation (with a of about 3 sec), while two components were observed in awake control studies (with values for of 3–6 and 20–40 s, respectively, Figure 16.11). This effect was not dependent on the intensity of the hypoxic stimulus nor on the CNS arousal state of the subject [23,24]. Sevoflurane, at 0.15 MAC, depressed the ventilatory response to multiple hypoxic pulses by about 30%. Analyzing the response using a two-compartment model revealed the presence of two components in control studies ( 3 and 100 s, respectively), while only one fast component was observed during sevoflurane inhalation [54,86]. The amplitude of both components was reduced by sevoflurane.
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Dahan et al. In awake studies, the ventilatory response to 20 min of isocapnic hypoxia shows a rapid return to pre-hypoxic baseline values without any undershoot. During administration of 0.15 MAC halothane, the relief of 20 min of hypoxia causes a large undershoot and irregular breathing followed by a very slow return to prehypoxic ventilation values [16]. We consider this an observation of STP inactivation. The post-hypoxic ventilatory pattern is now due to the loss of the hypoxic drive, the relief of HVD and BBF-related changes in brain tissue PCO2 with an initial low brain tissue PCO2 due to hypoxia followed by a gradual increase due to the return to normoxia. Upon square-wave increases and decreases in end-tidal PCO2, reduction by 60% of the time constant of the central chemoreflex loop is observed during 0.1 MAC halothane, isoflurane or sevoflurane inhalation. In awake humans, the time constant of the central chemoreflex loop (awake value about 2 min) is determined by BBF dynamics (CO2 wash-in into the brain compartment) and neuronal dynamics [45,49,60]. Since the anesthetics do not affect BBF at 0.1 MAC, the inactivation of STP is believed responsible for the reduction of the time constant of the central chemoreflex loop.
Interestingly, intravenous anesthetics, such as propofol, also inactivate STP. Like sevoflurane, propofol abolishes the slow component of the AHR [86]. However, in contrast to sevoflurane, the magnitude of the fast component of the AHR remains unaffected, indicating no direct effect of propofol at the CBs. The interaction of low-dose inhalational anesthetics with STP indicates that low-dose anesthetics have—apart from their peripheral site of action—a central effect site, independent of the arousal state of the subject [23,24]. It has been postulated [112] that STP is related to neural activity of respiratory neurons in the ponto-medullary brainstem, which results in local acidosis. The low pH then provides a strong stimulus to the adjacent local chemosensitive structures and consequently contributes to STP formation. Since low-dose inhalational anesthetics do not affect the central chemosensors [45,49,60], their effect on STP is best explained by a direct effect at the respiratory neurons in the brainstem. Another theory links the formation of STP to S-nitrosothiols (SNOs) such as S-nitrosogluthatione [113–115]. According to this theory, SNOs are produced in the brainstem neurons by neuronal nitric oxidase synthase (activated by afferents from peripheral chemoreceptors) and by erythrocyte deoxygenation (from reduced thiols). These SNOs then act at the NTS to cause a ventilatory response with STP-like characteristics. How anesthetics interact with SNO formation remains unknown at present but may be related to the altered ability of deoxygenated blood to produce SNOs and/or to a reduced afferent input to the NTS.
Influence of Inhalation Anesthetics V.
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Conclusions
Older (published before 1990) and more recent studies on the effect of inhalational anesthetics on CB-mediated responses have led to important observations: 1.
Inhalational anesthetics depress the CB response to hypoxia at even subanesthetic doses. This effect is variable among different agents as well as among the species in which these agents are tested. Recent studies have increased our knowledge of the molecular sites of action of the anesthetics in the CB and their impact on oxygen sensing. The role of ROS in anesthesia-induced depression of the hypoxic drive and the ability to reverse or prevent this by the administration of relatively simple antioxidants seem especially exciting and promising. It may open new routes toward inexpensive measures to treat and prevent anesthesiarelated respiratory events in postoperative patients. Whether a similar treatment is effective in anesthesia-induced discoordination of upper airway musculature needs further study. 2. Low-dose inhalational anesthetics have a central effect site of action causing the inactivation of short-term potentiation of breathing, and for halothane, the facilitation of the development of hypoxic ventilatory decline. These effects are independent of the arousal state of the subjects/patients. The consequences of these observations in animals and healthy volunteers are difficult to judge for postoperative patients. Although inhalational anesthetics may persist for several hours in the body after surgery, and although hypoxia is not uncommon in this period [116], various other influences determine the ventilatory pattern. These include genetic predisposition, age, obesity, gender and hormonal condition, underlying disease, upper airway function and patency, ventilatory pump function, additional oxygen inhalation, posture, pain and pain relief, tactile-, auditory- and visual-stimulation, surgery, stress, sedation, residual muscle relaxation, etc. [35]. To date, a causal relationship among the depression of hypoxic drive, the loss of STP, the facilitation of HVD development, and the occurrence of respiratory events in the postoperative period is at best speculative. References 1.
Gonzalez, C., Almaraz, L., Obeso, A. and Rigual, R., Carotid body chemoreceptors: From natural stimuli to sensory discharges, Physiol. Rev. 74, 829–898, 1994.
678 2. 3.
4.
5.
6.
7.
8.
9.
10. 11.
12. 13.
14.
15.
16.
Dahan et al. Prabhakar, N.R., Oxygen sensing by the carotid body chemoreceptors, J. Appl. Physiol. 88, 2287–2295, 2000. Vizek, M., Picket, C.K. and Weil, J.V., Biphasic ventilatory response of adult cats to sustained hypoxia has central origin, J. Appl. Physiol. 63, 1659–1664, 1987. Wang, Z.Z., Stensaas, L.J., Dinger, B.G. and Fidone, S.J., Nitric oxide mediates chemoreceptor inhibition in the cat carotid body, Neuroscience 65, 217–229, 1995. Campanucci, V.A., Fearon, I.M. and Nurse, C.A., A novel O2-sensing mechanism in rat glossopharyngeal neurones mediated by a halothaneinhibitable background Kþ conductance, J. Physiol. 548, 731–743, 2003. Cunningham, D.J.C., Robbins, P.A. and Wolff, C.B., Integration of respiratory responses to changes in alveolar pressures of CO2 and O2 and in arterial pH, in Handbook of Physiology, The Respiratory System, Volume II: Control of Breathing, Part 2, Fishman, A.P., Cherniack, N.S., Widdicombe, J.G. and Geiger, S.R., eds., Bethesda, American Physiological Society, pp. 1–528, 1986. Knill, R.L. and Gelb, A.W., Ventilatory responses to hypoxia and hypercapnia during halothane sedation and anesthesia, Anesthesiology 49, 244–251, 1978. Knill, R.L., Manninen, P.H. and Clement, J.L., Ventilation and chemoreflexes during enflurane sedation and anaesthesia in man, Can. Anesth. Soc. J. 26, 353– 360, 1979. Knill, R.L., Kieraszewicz, H.Y., Dodgson, B.G. and Clement, J.L., Chemical regulation of ventilation during isoflurane sedation and anaesthesia in humans, Can. Anesth. Soc. J. 30, 607–614, 1983. Knill, R.L. and Clement, J.L., Site of selective action of halothane on the peripheral chemoreflex pathway in humans, Anesthesiology 61, 121–126, 1984. Knill, R.L. and Clement, J.L., Ventilatory responses to acute metabolic acidemia in humans awake, sedated, and anesthetized with halothane, Anesthesiology 62, 745–53, 1985. Easton, P.A., Slykerman, L.J. and Anthonisen, N.R. Ventilatory response to sustained hypoxia in normal adults, J. Appl. Physiol. 64, 906–911, 1986. Easton, P.A., Slykerman, L.J. and Anthonisen, N.R., Recovery of the ventilatory response to hypoxia in normal adults, J. Appl. Physiol. 64, 521–528, 1988. Suzuki, A., Nishimura, M., Yamamoto, K., Miyamoto, K., Kishi, F. and Kawakami, Y., No effect of brain blood flow on ventilatory depression during sustained hypoxia, J. Appl. Physiol. 66, 1674–1678, 1989. Berkenbosch, A., Dahan, A., DeGoede, J. and Olievier, I.C.W., The ventilatory response to CO2 of the peripheral and central chemoreflex loop before and after sustained hypoxia in man, J. Physiol. (Lond.) 456, 71–83, 1992. Dahan, A., van den Elsen, M.J.L.J., Berkenbosch, A., DeGoede, J., Olievier, I.C.W. and van Kleef, J.W., Influence of a subanesthetic concentration of halothane on the ventilatory response to step changes into and out of sustained hypoxia in healthy volunteers, Anesthesiology 81, 850–859, 1994.
Influence of Inhalation Anesthetics 17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27. 28.
29.
30. 31.
679
Dahan, A., Ward, D.S., van den Elsen, M., Temp, J. and Berkenbosch, A., Influence of reduced carotid body drive during sustained hypoxia on hypoxic depression of ventilation in humans, J. Appl. Physiol. 81, 565–572, 1996. Nieuwenhuijs, D., Sarton, E., Teppema, L. and Dahan, A., Propofol for monitored anesthesia care: Implications on hypoxic control of cardiorespiratory responses, Anesthesiology 92, 46–54, 2000. Fatemian, M., Nieuwenhuijs, D.J.F., Teppema, L.J., Meinesz, S., van der Mey, A.G.L., Dahan, A. and Robbins, P.A., The respiratory response to carbon dioxide in humans with unilateral and bilateral resections of the carotid bodies, J. Physiol. (Lond.) 549, 965–973, 2003. Dahan, A., Sarton, E., van den Elsen, M., van Kleef, J., Teppema, L. and Berkenbosch, A., Ventilatory responses to hypoxia in humans: Influences of subanesthetic desflurane, Anesthesiology 85, 60–68, 1996. Berkenbosch, A., DeGoede, J., Ward, D.S., Olievier, C.N. and VanHartevelt, J., Dynamic responses of the peripheral chemoreflex loop to changes in end-tidal O2, J. Appl. Physiol. 71, 1123–1128, 1991. Kirby, T.P., Wraith, P.K., DeCort, S.C., Airlie, M.A.A., Hill, J.E., Carson, E.R., Flenley, D.C. and Warren, P.M., Modelling the dynamic ventilatory response to hypoxia in normal subjects, J. Theor. Biol. 166, 135– 147, 1994. Dahan, A., van den Elsen, M.J.L.J., Berkenbosch, A., DeGoede, J., Olievier, I.C.W. and van Kleef, J.W., Halothane affects ventilatory afterdischarge in humans, Br. J. Anaesth. 74, 544–548, 1995. Dahan, A., van Kleef, J., van den Elsen, M., Valk, R. and Berkenbosch, A., Slow ventilatory dynamics after isocapnic hypoxia and voluntary hyperventilation in humans: Effects of isoflurane, Br. J. Anaesth. 76, 373–381, 1996. Dahan, A., Berkenbosch, A., DeGoede, J., van den Elsen, M., Olievier, I., van Kleef, J., Influence of hypoxic duration and posthypoxic inspired O2 concentration on short term potentiation of breathing in humans, J. Physiol. (Lond.) 488, 803–813, 1995. Berkenbosch, A., DeGoede, J., Ward, D., Olievier, C.N. and Van Hartevelt, J., Dynamic response of peripheral chemoreflex loop to changes in end-tidal CO2, J. Appl. Physiol. 64, 1779–1785, 1988. Robbins, P.A., Hypoxic ventilatory decline: Site of action, J. Appl. Physiol. 79, 373–374, 1995. Ward, D.S., Berkenbosch, A., DeGoede, J. and Olievier, C.N., Dynamics of the ventilatory response to central hypoxia in cats, J. Appl. Physiol. 68, 1107–1113, 1990. Georgopoulos, D., Holtby, S.G., Berezanski, D. and Anthonisen, N.R., Aminophylline effects on ventilatory response to hypoxia and hyperoxia in normal adults, J. Appl. Physiol. 67, 1150–1156., 1989. Dahan, A. and Ward, D.S., Influence of i.v. midazolam on the ventilatory response to sustained hypoxia in man, Br. J. Anaesth. 66, 454–457, 1991. Tatsumi, K., Pickett, C.K. and Weil, J.V., Effects of haloperidol and domperidone on ventilatory roll off during sustained hypoxia in cats, J. Appl. Physiol. 72, 1945–1952, 1991.
680 32.
33.
34.
35.
36.
37.
38. 39.
40.
41.
42. 43.
44.
45.
46.
Dahan et al. Andronikou, S., Shirahata, M., Mokashi, A. and Lahiri, S., Carotid chemoreceptor and ventilatory responses to sustained hypoxia and hypercapnia in the cat, Respir. Physiol. 72, 361–374, 1988. Kimura, H., Tanaka, M., Nagano, K., Niijima, M., Masuyama, S., Mizoo, A., Uruma, T., Tatsumi, K., Kuriyama, T., Masuda, A., Kobayashi, T. and Honda, Y., Possible role of the carotid body responsible for hypoxic ventilatory decline in awake humans, in Physiology and Pharmacology of CardioRespiratory Control, Dahan, A., Teppema, L. and van Beek, H., eds., Dordrecht, Kluwer Academic Publishers, pp. 11–19, 1998. Long, W., Giesbrecht, G.G. and Anthonisen, N.R., Ventilatory response to moderate hypoxia in awake chemodenervated cats, J. Appl. Physiol. 74, 805–810, 1993. Nieuwenhuijs, D., Cardio-Respiratory Control in the Perioperative Patient: From Bench to Bedside, Ph.D. dissertation, Leiden University, Leiden, The Netherlands, 2002. Ang, R.C., Hoop, B. and Kazemi, H., Role of glutamate as the central neurotransmitter in the hypoxic ventilatory response, J. Appl. Physiol. 71, 1480–1487, 1992. Ohtake, P.J., Torres, P.J., Gozal, Y.M., Graff, G.R. and Gozal, D., NMDA receptors mediate peripheral chemoreceptor afferent input in the conscious rat, J. Appl. Physiol. 84, 853–861, 1998. Kazemi, H. and Hoop, B., Glutamic acid and g-aminobutyric acid neurotransmitters in central control of breathing, J. Appl. Physiol. 70, 1–7, 1991. Georgopoulos, D., Berezanski, D. and Anthonisen, N.R., Increased chemoreceptor output and ventilatory response to sustained hypoxia in normal adults, J. Appl. Physiol. 66, 1157–1163, 1989. Duffin, J., Triscott, A. and Whitwam, J.G., The effect of halothane and thiopentone on ventilatory responses mediated by the peripheral chemoreceptors in men, Br. J. Anaesth. 48, 975–980, 1976. Tusiewicz, K., Bryan, A.C. and Froese, A.B., Contribution of changing rib cage-diaphragm interactions to the ventilatory depression of halothane anesthesia, Anesthesiology 47, 327–337, 1977. Knill, R.L. and Gelb, A.W., Peripheral chemoreceptors during anesthesia: Are the watchdogs asleep? (editorial), Anesthesiology 57, 151–152, 1982. Temp, J.A., Henson, L.C. and Ward, D.S., Does a subanesthetic concentration of isoflurane blunt the ventilatory response to hypoxia? Anesthesiology 77, 1116–1124, 1992. Temp, J.A., Henson, L.C. and Ward, D.S., Effect of 0.1 MAC isoflurane on two tests of the hypoxic ventilatory response, Anesthesiology 78, 739–750, 1994. Dahan, A., van den Elsen, M., Berkenbosch, A., DeGoede, J., Olievier, I.C.W., van Kleef, J.W. and Bovill, J.G., Effects of subanesthetic halothane on the ventilatory responses to hypercapnia and acute hypoxia in healthy volunteers, Anesthesiology 80, 727–738, 1994. Young, C.H., Drummond, G.B. and Warren, P.M., Effect of a sub-anaesthetic concentration of halothane on the ventilatory response to sustained hypoxia in healthy humans, Br. J. Anaesth. 71, 642–647, 1993.
Influence of Inhalation Anesthetics 47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
681
Nagyova, B., Dorrington, K.L., Gill, E.W. and Robbins, P.A., Comparison of the effects of sub-hypnotic concentrations of propofol and halothane on the acute ventilatory response to hypoxia, Br. J. Anaesth. 75, 713–718, 1995. van den Elsen, M., Dahan, A., Berkenbosch, A., DeGoede, J., van Kleef, J. and Olievier, I.C.W., Does subanesthetic halothane affect the ventilatory response to acute isocapnic hypoxia in healthy volunteers? Anesthesiology 81, 860–867, 1994. van den Elsen, M., Dahan, A., DeGoede, J., Berkenbosch, A. and van Kleef, J., Influences of subanesthetic isoflurane on ventilatory control in humans, Anesthesiology 83, 478–490, 1995. Sollevi, A. and Lindahl, S.G.E., Hypoxic and hypercapnic ventilatory responses during isoflurane sedation and anaesthesia in women, Acta Anaesth. Scand. 39, 931–938, 1995. Foo, I.T., Martin, S.E., Fried, M.J., Douglas, N.J., Drummond, G.B. and Warren, P.M., Does sleep enhance the effect of subanesthetic isoflurane on hypoxic ventilation? Anesth. Analg. 81, 751–756, 1995. Foo, I.T., Martin, S.E., Lee, R.J., Drummond, G.B. and Warren, P.M., Interaction of the effects of domperidone and sub-anaesthetic concentrations of isoflurane on immediate and sustained hypoxic ventilatory response in humans, Br. J. Anaesth. 74, 134–141, 1995. Sarton, E., Dahan, A., Teppema, L., van den Elsen, M., Olofsen, E., Berkenbosch, A. and van Kleef, J., Acute pain and central nervous system arousal do not restore impaired hypoxic ventilatory response during sevoflurane sedation, Anesthesiology 85, 295–303, 1996. Sarton, E., van der Wal, M., Nieuwenhuijs, D., Teppema, L., Robotham, J.L. and Dahan, A., Sevoflurane-induced reduction of hypoxic drive is sexindependent, Anesthesiology 90, 1288–1293, 1999. Dahan, A., Nieuwenhuijs, D., Olofsen, E., Sarton, E., Romberg, R. and Teppema, L., Response surface modeling of alfentanil–sevoflurane interaction on cardiorespiratory control and bispectral index, Anesthesiology 94, 982–91, 2001. Pandit, J.J., Manning-Fox, J., Dorrington, K. and Robbins, P.A., Effects of subanaesthetic sevoflurane on ventilation. 2: Response to acute and sustained hypoxia in humans, Br. J. Anaesth. 83, 210–216, 1999. Teppema, L.J., Nieuwenhuijs, D., Sarton, E., Romberg, R., Olievier, C.N., Ward, D.S. and Dahan, A., Antioxidants prevent depression of the acute hypoxic ventilatory response by subanaesthetic doses of halothane, J. Physiol. (Lond.) 544, 931–938, 2002. Teppema, L.J., Romberg, R., Sarton, E. and Dahan, A., Antioxidants prevent depression of the acute hypoxic ventilatory response by subanaesthetic isoflurane, Anesthesiology, in press, 2005. Pandit, J.J., The variable effect of low-dose volatile anaesthetics on the acute ventilatory response to hypoxia in humans: A quantitative review, Anaesthesia 57, 632–643, 2002. van den Elsen, M., Sarton, E., Teppema, L., Berkenbosch, A. and Dahan, A., Influence of 0.1 minimum alveolar concentration of sevoflurane, desflurane and
682
61.
62. 63. 64.
65. 66. 67.
68.
69.
70.
71.
72. 73.
74. 75.
76.
Dahan et al. isoflurane on dynamic ventilatory response to hypercapnia in humans, Br. J. Anaesth. 80, 174–82, 1998. Nagyova, B., Dorrington, K.L., Poulin, M.J. and Robbins, P.A., Influence of 0.2 minimum alveolar concentration of enflurane on the ventilatory response to sustained hypoxia in humans, Br. J. Anaesth. 78, 707–713, 1997. Cheng, S.-C. and Brunner, E.A., Effects of anesthetic agents on synaptosomal GABA disposal, Anesthesiology 55, 34–40, 1981. Franks, N.P. and Lieb, W.R., Background Kþ channels: An important target for volatile anesthetics? Nat. Neurosci. 2, 395–396, 1999. Pattel, A.J., Honore´, E., Lesage, F., Fink, M., Romey, G. and Ladzunski, M., Inhalational anesthetics activate two-pore-domain background Kþ channels, Nat. Neurosci. 2, 422–426, 1999. Lo´pez-Barneo, J., Pardal, R. and Ortega-Sa´enz, P., Cellular mechanisms of oxygen sensing, Ann. Rev. Physiol. 63, 259–287, 2001. Perez-Garcia, M.T. and Lo´pez-Lo´pez, J.R., Are Kv channels the essence of O2 sensing? Circ. Res. 86, 490–491, 2000. Perez-Garcia, M.T., Lo´pez-Lo´pez, J.R., Riesco, A.M., Hoppe, U.C., Marban, E., Gonzalez, C. and Johns, D.C., Viral gene transfer of dominant-negative Kv4 construct suppresses an O2-sensitive Kþ current in chemoreceptor cells, J. Neurosci. 20, 5689–5695, 2000. Riesco-Fagundo, A.M., Perez-Garcia, M.T., Gonzalez, C. and Lo´pez-Lo´pez, J.R., O2 modulates large-conductance Ca2þ-dependent Kþ channels of rat chemoreceptor cells by a membrane-restricted and CO-sensitive mechanism, Circ. Res. 89, 430–436, 2001. Hatton, C.J. and Peers, C., Effects of cytochrome P-450 inhibitors on ionic currents in isolated rat type I carotid body cells, Am. J. Physiol. 271, C85–C92, 1996. Pardal, R. and Lopez-Barneo, J., Carotid body thin slices: Responses of glomus cells to hypoxia and Kþ-channel blockers, Respir. Physiol. Neurobiol. 132, 69–70, 2002. Gonzalez, C., Sanz-Alfayate, G., Agapito, M.T., Gomez-Nin˜o, A., Rocher, A. and Obeso, A., Significance of ROS in oxygen sensing in cell systems with sensitivity to physiological hypoxia, Respir. Physiol. Neurobiol. 132, 17–41, 2002. Weir, E.K., Hong, Z., Porter, V.A. and Reeve, H.L., Redox signaling in oxygen sensing by vessels, Respir. Physiol. Neurobiol. 132, 121–130, 2002. DeGroot, H. and Noll, T., Halothane hepatotoxicity: Relation between metabolic activation, hypoxia, covalent binding, lipid peroxidation and liver cell damage, Hepatology 3, 601–606, 1983. DeGroot, H. and Sies, H., Cytochrome P-450, reductive metabolism, and cell injury, Drug Metabol. Rev. 20, 275–284, 1989. Spracklin, D.K. and Kharasch, E.D., Human halothane reduction in vitro by cytochrome P450 2A6 and 3A4: Identification of low and high KM isoforms, Drug Metabol. Dis. 26, 605–607, 1998. Kharasch, E.D., Hankins, D.C., Fenstamaker, K. and Cox, K., Human halothane metabolism, lipid peroxidation, and cytochromes P4502A6 and P4503A4, Eur. J. Clin. Pharmacol. 55, 853–859, 2000.
Influence of Inhalation Anesthetics 77.
78.
79.
80. 81.
82.
83. 84.
84a.
85.
86.
87.
88.
89.
90.
683
Sato, N., Fum, K., Yuge, O., Tanaka, A. and Morio, M., Suppressive effect of vitamin E on lipid peroxidation in halothane-administered guinea pig liver, In Vivo 6, 503–505, 1992. Buckler, K.J., Williams, B.A. and Honore´, E., An oxygen-, acid- and anaesthetic-sensitive TASK-like background potassium channel in rat arterial chemoreceptor cells, J. Physiol. (Lond.) 525, 135–142, 2000. Sirois, J.E., Lei, Q., Talley, E.M., Lynch, III, C. and Bayliss, D.A., The TASK-1 two-pore domain Kþ channel is a molecular substrate for neuronal effects of inhalation anesthetics, J. Neurosci. 20, 6347–6354, 2000. Pattel, A.J. and Honore´, E., Properties and modulation of mammalian 2P domain Kþ channels, Trends Neurosci. 24, 339–346, 2001. Hanley, P.J., Ray, J., Brandt, U. and Daut, J., Halothane, isoflurane and sevoflurane inhibit NADH:ubiquinonen oxireductase (complex I) of cardiac mitochondria, J. Physiol. (Lond.) 544, 687–693, 2002. Waypa, G.B. and Schumaker, P.T., O2 sensing in hypoxic pulmonary vasoconstriction: The mitochondrial door re-opens, Respir. Physiol. Neurobiol. 132, 81–91, 2002. Baysal, B.E. and Taschner, P.E.M., Phenotypic dichotomy in mitochondrial complex II genetic disease, J. Mol. Med. 79, 495–503, 2001. Baysal, B.E., Ferrell, R.E., Willet-Bozick, J.E., Lawrence, C.E., Myssiorek, D., Bosch, A., van der Mey, A., Taschner, P.E.M., Rubinstein, W.S., Myers, E.N., Richard, C.W., Cornelisse, C.J., Devilee, P. and Devlin, B., Mutations in SDHD, a mitochondrial complex II gene, in hereditary paraganglioma, Science 287, 848–851, 2000. Dahan, A., Taschner, P.E.M., Jansen, J.C., van der Mey, A., Teppema, L.J. and Cornelisse, C.J., Carotid body tumors in humans caused by a mutation in the gene for succinate dehydrogenase D (SDHD), Adv. Exp. Med. Biol. 551, 71–76, 2004. Eng, C., Kiuru, M., Fernandez, M.J. and Aaltonen, L.A., A role for mitochondrial enzymes in inherited neoplasia and beyond, Nat. Rev. Cancer 3, 193–202, 2003. Nieuwenhuijs, D., Sarton, E., Teppema, L.J., Kruyt, E., Olievier, I., van Kleef, J. and Dahan, A., Respiratory sites of action of propofol—absence of depression of peripheral chemoreflex loop by low-dose propofol, Anesthesiology 95, 889–895, 2001. Weiskopf, R.B., Raymond, L.W. and Severinghaus, J.W., Effects of halothane on canine respiratory responses to hypoxia with and without hypercarbia, Anesthesiology 41, 350–360, 1974. Davies, R.O., Edwards, M.W. and Lahiri, S.L., Halothane depresses the response of carotid body chemoreceptors to hypoxia and hypercapnia in the cat, Anesthesiology 57, 153–159, 1982. Stuth, E.A.E., Dogas, Z., Krolo, M., Kampine, J.P., Hopp, F.A. and Zuperku, E.J., Effects of halothane on the phrenic nerve responses to carbon dioxide mediated by carotid body chemoreceptors in vagotomized dogs, Anesthesiology 87, 1440–1449, 1997. Ide, T., Sakurai, Y., Aono, M. and Nishino, T., Contribution of peripheral chemoreception to the depression of the hypoxic ventilatory response during halothane anesthesia in cats, Anesthesiology 90, 1084–1091, 1999.
684
Dahan et al.
91.
Dahan, A., Olofsen, E., Teppema, L., Sarton, E. and Olievier, C., Speed of onset and offset and mechanisms of ventilatory depression from sevoflurane, Anesthesiology 90, 1119–1128, 1999. Berkenbosch, A., DeGoede, J., Olievier, C.N. and Quanjer, P.H., Sites of action of halothane on respiratory pattern and ventilatory response to CO2 in cats, Anesthesiology 57, 389–398, 1982. van Dissel, J.T., Berkenbosch, A., Olievier, C.N., DeGoede, J. and Quanjer, P.H., Effects of halothane on the ventilatory response to hypoxia and hypercapnia in cats, Anesthesiology 62, 448–456, 1985. Koh, S.O. and Severinghaus, J.W., Effect of halothane on hypoxic and hypercapnic ventilatory responses of goats, Br. J. Anaesth. 65, 713–717, 1990. Stuth, E.A.E., Dogas, Z., Krolo, M., Kampine, J.P., Hopp, F.A. and Zuperku, E.J., Dose-dependent effects of halothane on the phrenic nerve responses to acute hypoxia in vagotomized dogs, Anesthesiology 87, 1428–1439, 1997. Ponte, J. and Sadler, C.L., Effect of halothane, enflurane and isoflurane on carotid body chemoreceptor activity in the rabbit and the cat, Br. J. Anaesth. 62, 33–40, 1989. Hirshman, C.A., McCullough, R.E., Cohen, P.J. and Weil, J.V., Depression of hypoxic ventilatory response by halothane, enflurane and isoflurane in dogs, Br. J. Anaesth. 49, 957–962, 1977. Chatterjee, I.B., Majumder, A.K., Nandi, B.K. and Subramanian, N., Synthesis and some major functions of vitamin C in animals, Ann. NY Acad. Sci. 258, 24–47, 1975. von Euler, C., Brain stem mechanisms for generation and control of breathing pattern, in Handbook of Physiology, Volume II, Control of Breathing, Part I, Fishman, A.P., Cherniack, N.S., Widdicombe, J.G. and Geiger, S.R., eds., Bethesda, American Physiological Society, pp. 1–68, 1986. Orem, J. and Trotter, R.H., Behavioral control of breathing, News Physiol. Sci. 9, 228–231, 1994. Nishino, T. and Kochi, T., Effects of surgical stimulation on the apneic threshold for carbon dioxide during anaesthesia with sevoflurane, Br. J. Anaesth. 73, 583–586, 1994. Lam, A.M., Clement, J.L. and Knill, R.L., Surgical stimulation does not enhance ventilatory chemoreflexes during enflurane anesthesia in man, Can. Anaesth. Soc. J. 27, 22–28, 1980. Rosenberg, M., Tobias, B., Bourke, D.L. and Kumat, V., Respiratory responses to surgical stimulation during enflurane anesthesia, Anesthesiology 52, 163–165, 1983. Sarton, E., Dahan, A., Teppema, L., Berkenbosch, A., van den Elsen, M. and van Kleef, J., Influence of acute pain induced by activation of cutaneous nociceptors on ventilatory control, Anesthesiology 87, 289–296, 1997. Sirois, J.E., Lynch, III, C. and Baylis, D.A., Convergent and reciprocal modulation of a leak Kþ current and Ih by an inhalational anaesthetic and neurotransmitters in rat brainstem motoneurones, J. Physiol. (Lond.) 541, 717–729, 2002.
92.
93.
94. 95.
96.
97.
98.
99.
100. 101.
102.
103.
104.
105.
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106. Eldridge, F.L., Kiley, J.P. and Paydarfar, D., Dynamics of medullary hydrogen ion and respiratory neurones to square-wave change of arterial carbon dioxide in man, J. Physiol. (Lond.) 385, 627–642, 1987. 107. Younes, M., The physiological basis of central apnea and period breathing, Curr. Pulm. 10, 265–326, 1989. 108. Eldridge, F.L., Subthreshold central neural respiratory activity and after discharge, Respir. Physiol. 39, 327–343, 1980. 109. Teppema, L.J., Vis, A., Evers, J.A.M. and Folgering, H.T., Dynamics of brain extracellular fluid pH and phrenic nerve activity in cats after end-tidal CO2 forcing, Respir. Physiol. 50, 359–380, 1982. 110. Wilkins, C.J., Reed, P.N. and Aikenhead, A.R., Hypoxaemia after inhalation of 50% nitrous oxide and oxygen, Br. J. Anaesth. 63, 346–347, 1989. 111. Nortwood, D., Sapsford, D.J., Jones, J.G., Griffith, D. and Wilkins, C., Nitrous oxide sedation causes post-hyperventilation apnea, Br. J. Anaesth. 67, 7–12, 1991. 112. Santiago, T.V. and Edelman, N.H., Brain blood flow and control of breathing, in Handbook of Physiology, Volume II, Control of Breathing, Part I, Fishman, A.P., Cherniack, N.S., Widdicombe, J.G. and Geiger, S.R., eds., Bethesda, American Physiological Society, pp. 163–179, 1986. 113. Lipton, A.J., Johnson, M.A., MacDonald, T., Lieberman, M.W., Gozal, D. and Gaston, B., S-Nitrosothiols signal the ventilatory response to hypoxia, Nature 413, 171–174, 2001. 114. Gozal, D., Gaston, B., Lipton, A.J., Johnson, M.A., MacDonald, T. and Lieberman, M.W., The ventilatory response to hypoxia: Reply, Nature 419, 686, 2002. 115. Lipton, S.A., Nitric oxide and respiration, Nature 413, 118–121, 2001. 116. Catley, D.M., Thornton, C., Jordan, C., Lehane, J.R., Royston, D. and Jones, J.G., Pronounced episodic oxygen desaturation in the postoperative period, Anesthesiology 63, 20–28, 1985.
17 General Anesthesia and Respiratory Mechanics
DAVID O. WARNER Mayo Clinic College of Medicine Rochester, Minnesota
I.
Introduction
The study of respiratory mechanics concerns the motion of structures that control the movement of air into and out of the lungs. One set of structures controls intrathoracic pressure, which drives gas flow; these structures comprise the chest wall. Another system maintains the patency of the extrathoracic airway proximal to the glottis; the components of this system comprise the upper airway. This chapter will describe the functional consequences of anesthetic-induced alterations in the activation of skeletal muscles that control the chest wall and upper airway (often referred to as respiratory muscles). Anesthetics also have important effects on smooth muscle lining the airways, which also regulates gas flow within the lungs. This topic is reviewed elsewhere [1] and is beyond the scope of this chapter. Historically, simple observation of external respiratory system motion provided a powerful tool to guide the proper administration of anesthesia [2]. More recently, alterations in the control of respiratory muscles that lead to changes in respiratory mechanics have been invoked to explain much of the impairment of gas exchange observed during and after clinical anesthesia. 687
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Thus, anesthetic-induced changes in respiratory muscle activation and respiratory mechanics can have profound clinical consequences. II.
Normal Function of the Respiratory Pump
A detailed description of the components and function of the respiratory pump is available in other texts [3,4]. Here summarized are the features necessary to further discuss anesthetic effects. The two major components of the respiratory system germane to this discussion of mechanics include the chest wall, which generates the changes in intrathoracic pressure that produce gas flow, and the upper airways, which conduct gas from the atmosphere to the trachea. A. Chest Wall Components
The diaphragm separates the contents of the thorax and abdomen. Anatomically, the muscle consists of crural and costal portions, which both insert into a central tendon [5]. The two parts may have different functions under some circumstances [6,7], although they both participate in most respiratory behaviors. The human diaphragm can be considered an elliptical cylindroid capped by an irregular dome [5] (Figure 17.1). The cylindrical portion is adjacent to the inner surface of the rib cage and so conforms to its shape in this area of apposition [8]. When the diaphragm contracts, it causes a caudad motion of the dome [9,10]. In recumbent postures, the axial displacement of dependent regions (e.g., posterior in supine subject) exceeds that of the nondependent regions [9,11–13]. Isolated contraction of the diaphragm also affects other chest wall structures such as the rib cage by direct mechanical actions via its insertions, and by changing abdominal and thoracic pressures; the significance of these effects depends on posture, lung volume, and the activity of other chest wall muscles [6,7,14–16]. For example, diaphragm contraction expands the lower rib cage by two mechanisms: increases in abdominal pressure, transmitted to the rib cage through the area of apposition, and elevation of the lower ribs by the force exerted by the diaphragm. The rest of the thorax is surrounded by the rib cage, which provides rigidity and a point of attachment for all the chest wall muscles (Figure 17.2). The actions of the intrinsic muscles of the rib cage, such as the intercostal muscles, are complex and still not completely understood [17]. In general, the muscles of the more superficial layers (e.g., parasternal intercostals) elevate the ribs and thus have an inspiratory action, whereas the deeper layers tend to depress the ribs and promote expiration [18,19]. However, the net effect of any individual muscle depends on several factors such as lung
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Ribs
Pes
Diaphragm
Pga Area of apposition
Figure 17.1 Coronal section of the thorax, showing the dome of the diaphragm, the area of apposition where the diaphragm is adjacent to the inner surface of the rib cage, and the regions where esophageal (Pes) and gastric (Pga) pressures are measured to quantify chest wall function (Data from Ref. 4).
volume [18,20,21] and the activity of adjacent muscles [18]. These muscles also serve a stabilizing function, preventing rib cage distortion. For example, isolated diaphragmatic contraction decreases intrathoracic pressure, which causes retraction of the upper rib cage [14,22,23]. Normally, contraction of intercostal muscles prevents this retraction [16,24]. Other muscles such as the scalenes and sternocleidomastoids also insert onto the bony rib cage and have respiratory actions to elevate the ribs and assist inspiration. Although previously referred to as accessory muscles of respiration, perhaps implying that they are used only infrequently, they may be active during even quiet breathing [25,26]. Finally, abdominal muscles also have important respiratory actions. Contraction of these muscles raises intra-abdominal pressure, which tends to move the diaphragm cephalad and thus decrease lung volume [27]. However, this expiratory action may be opposed by direct actions of abdominal muscles via insertions on the rib cage [28] and by increased abdominal pressure that may act, via the area of apposition between diaphragm and rib cage, to expand the lower rib cage [29].
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Scalenus anterior muscle
External intercostal muscle
Anterior intercostal membrane
Scalenus medius muscle
Parasternal intercostal muscle
Internal intercostal muscle
Figure 17.2 Anatomy of the rib cage muscles. The external intercostal muscles and anterior scalene muscles have been removed from the left side of the chest to show the internal intercostal and medial scalene muscles, respectively (redrawn from Ref. 210).
Normal Chest Wall Mechanics
The respiratory muscles move the chest wall. Due to the complexities of their anatomic arrangement, and the fact that these muscles serve other functions such as the maintenance of posture (often simultaneously with breathing), it is difficult to assign specific actions to individual respiratory muscles. Rather, the net effect of this activity is estimated by measuring (1) the shape and motion of the chest wall, and (2) consequent changes in lung volume and respiratory system pressures. The rib cage expands and the diaphragm descends with inspiration. Accommodation of diaphragmatic descent requires an expansion of the abdomen because its contents are relatively incompressible. Changes in lung volume can be accurately estimated from measurements of the external dimensions of the rib cage and the abdomen [30]. These measurements can be readily obtained by using various devices that measure anteroposterior diameters, circumferences, or cross-sectional areas of the rib cage and abdomen (Figure 17.3). Furthermore, the tidal volume can be partitioned into a volume displaced by the rib cage and
Bands
Rib cage area (volume)
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"Isovolume" lines
Tidal volume
Abdominal area (volume)
Figure 17.3 Schematic representation of system used to describe chest wall motion using changes in external dimensions of the rib cage and abdomen first proposed by [30]. Axes in graph depict the changes in cross-sectional area contained within the bands; with calibration, they also represent volumes displaced by the respiratory motion of the rib cage and diaphragm-abdomen compartments. Solid line in graph shows typical pattern during quiet breathing. Dashed lines represent isovolume lines; all points that fall on a given isovolume line represent chest wall configurations that result in the same lung volume.
a volume displaced by the diaphragm-abdomen, providing a convenient description of the pattern of chest wall motion. These volumes do not precisely correspond to volumes swept by the anatomic surfaces of the rib cage and the diaphragm, because the rib cage and diaphragm are mechanically coupled at the area of apposition and the ventral abdominal wall (Figure 17.1) [31,32]. However, this is a useful method to describe chest wall behavior. Pressures in the thorax (usually measured in the esophagus) and the abdomen (usually measured in the stomach) can be used to calculate transdiaphragmatic pressure, an estimate of the effective force generated by diaphragm contraction (Figure 17.1). Various indices of diaphragm and rib cage function have been calculated using combinations of these pressures [33,34]. However, attributing changes in these pressures to the actions of specific muscles, such as the diaphragm, must be done cautiously when other respiratory muscles are also active. For example, postoperative measurements of transdiaphragmatic pressure do not directly reflect diaphragm activity if abdominal muscle activity, which also influences gastric pressure, is also present [35]. Measurements of pressures, and simultaneous measurements of displacements such as changes in lung volume, can also be used to describe the elastic properties of the chest wall (in the absence of respiratory muscle activity) through familiar means such as the relationship between airway pressure and lung volume (a ‘pressure-volume curve’) [36].
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One important parameter of pulmonary mechanics is the functional residual capacity (FRC), the volume of gas in the lung at the end of expiration. In general, decreases in FRC are associated with an impaired ability of the lung to exchange gas. During quiet breathing in adults, the FRC is determined largely by the passive mechanical properties of the chest wall; namely the balance between the inward elastic recoil of the lung and the outward elastic recoil of the thorax [36]. Tonic activity in muscles with inspiratory actions such as the diaphragm has been thought to help maintain FRC, but the presence of such activity is controversial [12,37–42]. Younger children, whose chest wall is relatively more compliant, rely on mechanisms such as end-expiratory glottic constriction to maintain FRC [43–45], but this mechanism does not operate in normal adults [36]. B. Upper Airway Components
Many muscles and other structures surround the upper airway and determine its caliber [4,46]. We here concentrate on those structures most likely to influence pharyngeal and laryngeal patency during anesthesia. Palate and pharynx
The soft palate extends from the posterior border of the hard palate into the oropharynx. Its position, which determines gas flow through the mouth or nose, is regulated by five muscles: the tensor and levator veli palatini, musculus uvulae, palatoglossus, and palatopharyngeus. Pharyngeal caliber is narrowed by three overlapping muscles composing the lateral and posterior walls of the pharynx: the superior, middle, and inferior constrictors. Anterior and lateral muscles dilate the pharynx. The tongue forms the anterior wall of the oropharynx, and the hyoid bone and associated muscles form the anterior wall of the hypopharynx. The intrinsic muscles of the tongue primarily control shape. The position of the tongue is controlled by several extrinsic muscles connecting the tongue with surrounding structures. The genioglossus causes tongue protrusion, and is primarily responsible for anterior motion of the posterior part of the tongue to enlarge the oropharynx [47]. The genioglossus also acts with the hypoglossus to depress the tongue, while two muscles, the styloglossus and the palatoglossus, elevate the tongue. Several muscles attached to the hyoid bone may control pharyngeal patency. The geniohyoid, mylohyoid, stylohyoid, and digastric muscles attach to the superior hyoid and elevate it; the omohyoid, thyrohyoid, and sternohyoid muscles attach to the inferior hyoid and lower it. The coordinated activity of these muscles increases pharyngeal caliber and minimizes the descent of the larynx during inspiration [48,49].
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Larynx
Extrinsic muscles control laryngeal position via attachments to outside structures such as the sternum and include many of the muscles previously described as controlling hyoid position. Intrinsic laryngeal muscles control patency of the laryngeal inlet and the position of the vocal cords. The laryngeal inlet is narrowed by contraction of the transverse arytenoids, oblique arytenoids, and lateral cricothyroids. The posterior cricoarytenoids, which abduct the vocal cords, are the primary muscles controlling the laryngeal aperture in adults during resting breathing [50,51]. Other muscles with known respiration-related activity in humans include the cricothyroids [52], which tense the vocal cords, and the thyroarytenoids [53], which relax and adduct the vocal cords. Normal Upper Airway Mechanics
Description of upper airway mechanics is complicated by the fact that the upper airway actually serves several functions in addition to the maintenance of gas flow to the lungs, including deglutition and speech. Also, assessment of upper airway mechanics in general is more difficult compared with chest wall mechanics, because of difficulties in measuring the appropriate pressures and assessing the motion of upper airway structures. However, recent appreciation of the importance of upper airway physiology to diseases such as obstructive sleep apnea has spurred considerable advances in the assessment of upper airway function. Imaging modalities such as magnetic resonance imaging and fiberoptic pharyngoscopy have provided structural information, and assessment of pressure flow relationships by using catheters positioned at appropriate levels of the upper airway permit measurements of dynamic function. Although the specific action of individual muscles is not easily defined, there is little doubt that their combined effect is critical in maintaining airway patency, especially during inspiration when contraction of the chest wall muscles generates negative pressure in the upper airway (Figure 17.4) [54]. The genioglossus, stylopharyngeus, styloglossus, posterior cricoarytenoid, cricothyroid muscles, and perhaps others may all contribute to dilating the pharynx and larynx during inspiration [50–52,55–57]. The glottis widens during inspiration and narrows during expiration in human subjects, primarily due to phasic activation of the posterior cricoarytenoid muscle, a vocal cord abductor [58–60]. As is the case for many upper airway muscles, activation of this muscle precedes activation of chest wall muscles and, consequently, precedes inspiratory flow. This pattern of activation may minimize airflow resistance by opening the vocal cords prior to inspiration. The activity decreases during expiration, and the resulting glottic narrowing may serve to retard and control expiratory flow [59,61]. Phasic expiratory
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Airway AP diameter (% control)
100 80 60 40 Active 20 Passive 0 −15
−10
−5
0
Mouth pressure (cmH2O)
Figure 17.4 Relationship between anterioposterior upper airway diameter (measured by fluoroscopy) and mouth pressure in six supine subjects when mouth pressure is decreased by graded inspiratory effort against an occluded airway (active upper airway muscles) or negative pressure generated by an external source (passive upper airway muscles). Note that airway size is maintained despite negative pressures when upper airway muscles are active, and that the upper airway is markedly narrowed by relatively modest pressures in the absence of activity, demonstrating the important of upper airway muscle activity in maintaining upper airway patency (redrawn from Ref. 54).
activity in vocal cord adductors such as the thyroarytenoids may also serve to control expiratory flow [53,62]. The extrinsic laryngeal muscles can also dilate the upper airway [48,49]; however, little information is available regarding the actions of these muscles in humans. It is likely that the coordinated activity of all these muscles is necessary to maintain normal upper airway caliber. III.
Effects of Anesthesia on Chest Wall Mechanics
General anesthesia profoundly affects chest wall mechanics during spontaneous breathing by altering the activation of respiratory muscles. We consider here these effects by looking first at the effects of general anesthesia on end-expiratory chest wall configuration (i.e., at FRC), an important determinant of gas exchange. We then examine anesthetic effects on the motion of the chest wall. In clinical practice, the functions of the respiratory
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muscles during anesthesia are often supplemented or entirely supplanted by positive pressure ventilation, which itself profoundly alters chest wall mechanics, a situation that is not further discussed here [63]. Before proceeding, it is important to recognize a fundamental challenge in the study of this area—respiratory muscle behavior differs significantly among species. The chest wall of quadrupeds differs from that of humans in both function and form. For example, during quiet breathing in dogs, phasic activity is prominent in muscles with expiratory actions, such as the abdominal and triangularis sterni muscles [23,64–70]. Thus, expiration is an active process in the dog even while at rest, unlike humans [12,40]. Consequently, the lung volume at the end of expiration is less than the lung volume when all muscles are relaxed. Dogs then initiate inspiration by relaxing their expiratory muscles and allowing lung volume to increase passively; activation of the diaphragm and other inspiratory muscles completes the inspiration. The opposite sequence occurs during expiration (Figure 17.5). This active expiratory activity may be responsible for a significant portion of the tidal volume [22,23,69–72]. Active expiration is generally observed in normal awake human subjects only during stimulated breathing, such as occurs during exercise [73]. The reason why these animals use such a markedly different strategy of breathing compared with humans is not known, but may involve adaptation to the typical posture maintained by each species; homo sapiens is the only species (including other primates) that spends a significant amount of time in the upright position. In addition to these differences in awake behavior, there are significant differences among species in the response of respiratory mechanics to anesthetic drugs. For example, in dogs the pattern of activation during wakefulness is maintained with pentobarbital anesthesia. However, parasternal intercostal activity during quiet breathing is maintained during anesthesia with the volatile agents, unlike findings in most human subjects [74]. More strikingly, the volatile anesthetics decrease or abolish phasic expiratory muscle activity in dogs and cats, in marked distinction to their stimulating effects on such activity in many human subjects [75]. As a result, anesthesia with the volatile agents in dogs either has no effect or may actually increase the FRC [74,76], as phasic expiratory muscle activity that reduces end-expiratory lung volume is lost with the induction of anesthesia. Because of these significant species differences in the behavior of the respiratory muscles in situ, subsequent discussion concentrates only on findings from human studies. A. Anesthetic Effects on Functional Residual Capacity
In most recumbent human subjects, the induction of general anesthesia reduces the FRC [63,77]. This decrease occurs rapidly after anesthetic induction and does not change over time (Figure 17.6) [12,42,78–82]. The
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HALOTHANE
Transverse abdominis External oblique Triangularis sterni Parasternal intercostal Crural diaphragm Costal diaphragm Esophageal 5 cmH O 2 pressure Gastric pressure
5 cmH2O 1s
Figure 17.5 Pattern of respiratory muscle electromyogram activity and chest wall pressures during pentobarbital and halothane anesthesia in a dog lying supine. The pattern of breathing during pentobarbital anesthesia resembles that present while dogs are awake, with prominent phasic activity in rib cage (triangularis sterni) and abdominal (transversus abdominis and external oblique) muscles with expiratory actions. Halothane anesthesia abolishes this activity, whereas phasic inspiratory activity in parasternal intercostal muscles is maintained. This significantly changes the pressures generated in both the thorax and abdomen. This effect of halothane is markedly different from that observed in human subjects (compare with Figure 17.9) (Data from Ref. 74).
addition of muscular paralysis does not further decrease the FRC [82]. Posture appears to be an important factor, as the FRC does not decrease if anesthesia is induced in the sitting position [83]. Decreases in FRC are exaggerated in the obese [84–86]. Most anesthetic drugs, including thiopental, methoxyflurane, propofol, halothane, and isoflurane, decrease the FRC; however, the FRC is not affected by ketamine or, under some circumstances, methohexital anesthesia [87–89]. Although the ultimate causes for the decrease in FRC remain largely unknown, recent studies have provided new insight into proximate mechanisms. Before discussing mechanisms involving the respiratory muscles, two other possible mechanisms should be mentioned. First, under some circumstances, increases in intrathoracic blood volume may
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Spirometer volume
300 ml Time Thiopental
Succinylcholine
Figure 17.6 Spirogram during induction of anesthesia in a human subject with thiopental, demonstrating an immediate decrease in functional residual capacity that is not further affected by muscular paralysis. The spirogram has been modified to account for oxygen uptake (redrawn from Ref. 78).
displace gas and reduce FRC, although this finding is controversial [12,42,90–92]. The cardiovascular effects of anesthetic drugs presumably cause these changes in blood volume. Second, any increases in lung elastic recoil would tend to decrease the FRC, and recoil is indeed increased by anesthesia [82]. However, this increase is now thought to be secondary to breathing at low lung volumes, and not a primary effect of anesthetics on lung parenchyma. Rather, current evidence suggests that anesthetic-induced reductions in FRC are largely produced by alterations in the control of the respiratory (and perhaps other) muscles [63]. It is convenient to consider the possible mechanisms of effects of anesthesia on FRC in terms of the displacement of the two primary components of the chest wall: the rib cage and the diaphragm. Inward Rib Cage Displacement
Although initial studies suggested otherwise [93,94], it is now apparent that internal rib cage cross-sectional area at end expiration consistently decreases during general anesthesia, and is at least partly responsible for the observed decreases in FRC [12,42,90,91,95,96]. At least three mechanisms have been advanced to explain decreases in end-expiratory rib cage dimensions produced by anesthesia (Figure 17.7). Abolition of tonic inspiratory muscle activity
Although the existence and importance of tonic activity in inspiratory agonists has been controversial, recent studies have failed to find such activity in supine human subjects, suggesting that constriction of the rib cage caused by anesthesia does not result from abolition of this activity
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Rib cage
Cephalad
Thorax
Abdomen Awake Anesthetized
Spine Plane used to define dependent diaphragm motion
Figure 17.7 Diagrammatic representation of a mid-sagittal section of the chest wall of subjects lying supine, showing the position of chest wall structures at functional residual capacity during quiet breathing while awake (solid lines) and while anesthetized with halothane (dashed lines). Halothane causes an increase in spinal curvature, an inward and caudad motion of the anterior rib cage, and changes diaphragmatic shape. Also shown is the plane used to define the axial motion of dependent (posterior) portion of the diaphragm (see Figure 17.17) (Data from Ref. 194).
[32,37–40,42]. If tonic activity in chest wall muscles is important to the maintenance of FRC (see above), abolition of such activity by anesthesia with muscular paralysis alone (without anesthesia) should reduce the FRC. Consistent with this idea, some studies have found that partial neuromuscular blockade decreases the FRC in seated awake subjects [37,97]. However, other studies have found little change [98,99], and it is possible that changes in the volume of blood in the thorax, not changes in chest wall dimensions, were responsible for observed alterations in the FRC. The FRC does not change with partial paralysis in supine subjects [100]. Changes in thoracic curvature
Halothane anesthesia with spontaneous breathing increases the curvature of the thoracic spine in the sagittal plane (Figure 17.7) [12,42], and the addition of paralysis does not further change thoracic curvature. This increase in curvature may contribute to rib cage constriction [101]. The mechanism producing changes in spinal curvature is unknown. If thoracic back muscles exhibit tonic activity while subjects are awake in the supine position, anesthesia could suppress this activity; however, it is not known if such activity exists. Expiratory activity in the rectus abdominis muscle could also flex the spine; however, such activity is rarely observed during breathing maneuvers [102].
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Phasic activity of expiratory muscles
In most subjects, general anesthesia produces phasic expiratory activity in abdominal and rib cage muscles [42,103–106]. When present, this activity has a clear expiratory effect on the rib cage, producing end-expiratory constriction of the rib cage that is abolished by paralysis [42]. The source of this action is most likely intrinsic rib cage muscles such as the internal intercostal or the triangularis sterni muscles [28,105,107,108]. However, this phasic expiratory activity is not necessary for a decrease in end-expiratory rib cage dimensions, as it also occurs when expiratory muscles are paralyzed [42]. Also, decreases in FRC produced by anesthesia and paralysis are similar in magnitude to those produced by anesthesia with spontaneous breathing [42,80,82]. Thus when present, expiratory muscle activity may contribute to decreases in FRC, but such decreases are still observed even when it is absent. Cephalad Diaphragm Displacement
Although earlier studies suggested that a cephalad (headward) motion of the end-expiratory position of the diaphragm contributed to reductions in the FRC produced by anesthesia [11,90], subsequent work has consistently failed to find a significant contribution, either during spontaneous breathing or paralysis with mechanical ventilation [9,12,13,42,91,109]. Rather, anesthesia consistently changes the shape of the diaphragm in subjects lying supine, with cephalad motion of the most dependent regions and a caudad motion of the most nondependent regions (Figure 17.8). Several factors may be responsible for these changes in diaphragm shape. Abolition of tonic diaphragm activity
The observed change in diaphragmatic shape may be related to changes in the neural activation of the diaphragm. Changes in diaphragm position with anesthesia have been attributed to loss of tonic diaphragmatic activity [11,41,110]. However, like tonic rib cage muscle activity, the presence of such tone in awake subjects lying supine is at best controversial [40], and has not been observed in direct measurements using intramuscular EMG electrodes in supine human subjects [12,42]. Changes in the position of diaphragm insertions
Alterations in the shape of the diaphragm may also be related to motions of its insertions on the thoracoabdominal wall. Because the anterior diaphragm inserts near the costal margin, motion of the anterior rib cage may also affect the diaphragm. For example, changes in the end-expiratory position of the anterior diaphragm with the induction of anesthesia appear to be closely related to motion of the rib cage [12]. Also, because the posterior portion of the diaphragm inserts on vertebral bodies, the increase
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Figure 17.8 Changes in the average end-expiratory axial position of the diaphragm as a function of vertical (anteroposterior) distance from the most dependent regions, measured using fast volumetric CT scanning, with the induction of 1.2 MAC halothane anesthesia in six supine human subjects. Note that the posterior (dependent) regions consistently moved cephalad (headward) and the anterior (non-dependent) regions moved caudad in five subjects. The net volume displaced by the diaphragm at end-expiration during the transition from wakefulness to anesthesia (28 68 ml, mean S.D.) did not differ significantly from 0, indicating that changes in diaphragm position do not consistently contribute to reductions in FRC caused by anesthesia (Data from Ref. 12).
in spinal curvature noted above may tend to move the posterior portion of the diaphragm cephalad (Figure 17.7). Phasic activity of expiratory muscles
Finally, expiratory activity in the abdominal muscles should increase pressure within the abdomen and tend to displace the diaphragm cephalad. The transversus abdominis, which exhibits phasic expiratory activity in many subjects during anesthesia [42,106], is particularly efficient at increasing abdominal pressure [102]. However, pharmacologic paralysis of this activity in fact produces a small but significant cephalad motion of the diaphragm [42]. Thus, it appears that phasic activity by the transversus abdominis muscle during halothane anesthesia does not have a significant mechanical effect on diaphragm position. The mechanism of this small cephalad motion of the diaphragm produced by paralysis is unclear. Diaphragm position would be affected by changes in intrathoracic or abdominal pressures; however, paralysis does not produce significant changes in either esophageal or gastric pressures measured at end-expiration compared with those measured during spontaneous breathing [42].
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Summary of Anesthetic Effects on FRC
It is now clear there are many mechanisms responsible for anesthesiainduced decreases in FRC, and different mechanisms may combine in individual patients depending on factors such as gender, body habitus, and the presence of respiratory system disease. In general, an inward motion of the rib cage contributes to reductions in FRC. Although anesthesia produces a change in diaphragm shape, net changes in diaphragm position do not consistently contribute to changes in FRC. Under some circumstances, changes in intrathoracic blood volume may also contribute to anesthesia-induced changes in the FRC. B. Anesthetic Effects on Chest Wall Motion
Anesthetic drugs change the activation of respiratory muscles during spontaneous breathing, and thus can affect the chest wall motion. We first discuss the effects of anesthesia on the pattern of respiratory muscle activation, then the consequences to the motion of the two primary components of the chest wall: the rib cage and the diaphragm [111]. Most experimental work in human subjects has been done using halothane as a prototypical volatile anesthetic, and thus much of the information reviewed in this section concerns this agent. Although not systematically studied, it is likely that the effects of halothane on respiratory muscle activation are similar to those of other potent volatile agents, based on animal studies and on studies of overall ventilatory control. Respiratory Muscle Activation
Anesthetic drugs have profound effects on overall respiratory drive and timing, as described in chapters 15 and 16. However, these effects are not distributed equally among different groups of respiratory muscles. Respiratory muscles other than the diaphragm
When humans breathe quietly in the supine position, there is phasic inspiratory activity in the diaphragm, in intercostal muscles with inspiratory actions such as the parasternal intercostal muscles, and in some accessory muscles such as the scalenes [38,40,105,107,112–114]. Muscles with expiratory actions, such as the transversus abdominis, are normally not active [40,42,106]. With the induction of anesthesia, two major changes in this pattern have been noted in several studies. First, there is an inhibition (or abolition) of activity in parasternal intercostal muscles [12,94,106,115], and second, phasic expiratory activity develops in some abdominal muscles (preferentially the transversis abdominis muscle, the innermost layer) [12,103,104,106]. Expiratory muscle activity in intrinsic rib cage muscles, such as the internal intercostals, can also be present during anesthesia
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Figure 17.9 Pattern of breathing in one male subject before (left panel) and after (right panel) the induction of halothane anesthesia, showing electromyogram activity measured with fine-wire percutaneous electrodes and the cross-sectional areas of the rib cage and abdomen during quiet breathing. Open and closed circles denote the beginning and end of inspiratory gas flow, respectively. The diaphragm electrode has been placed through the lateral rib cage intercostal muscles into the diaphragm at the area of apposition (see Figure 17.1). Halothane, 1.2 MAC end-tidal concentration, increases respiratory rate, abolishes activity in the parasternal intercostal muscle, and produces phasic expiratory activity in the transversus abdominis muscle. Phasic expiratory activity is also observed in the diaphragm electrode, probably representing activity in the adjacent internal intercostal muscles. Changes in rib cage and abdominal cross-sectional areas during breathing are proportionally decreased by halothane anesthesia, with relative preservation of rib cage expansion, despite the lack of parasternal intercostal activity (Data from Ref. 12).
(Figure 17.9) [12,106]. However, it now appears that the pattern of respiratory muscle use can differ considerably among humans anesthetized with a given agent. In a small study of six subjects, Warner et al. [42] found that differences in respiratory muscle activation were associated with gender. Male subjects given approximately 1.2 MAC end-tidal halothane in oxygen exhibited prominent phasic activity in expiratory muscle during quiet breathing, with no parasternal intercostal muscle activity and scalene muscle activity in only one of three subjects. In contrast, expiratory muscle activity was absent in three anesthetized female subjects during quiet breathing, whereas parasternal intercostal and scalene muscle activities were preserved (Figure 17.10). All subjects developed phasic expiratory activity in the transversus abdominis and phasic inspiratory activity in the scalene
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Figure 17.10 EMG recording from one female subject while awake (left panel), during the induction of halothane anesthesia when end-tidal halothane concentration was equivalent to approximately 0.4 MAC (middle panel), and after achieving a stable 1 MAC end-tidal concentration of halothane (right panel). Open and closed circles denote the beginning and end of inspiratory gas flow, respectively. Note that there is only transient activation of the transversus abdominis muscle during induction, and that parasternal intercostal and scalene activities are preserved during anesthesia. This pattern is in marked contrast to that observed in male subjects (see Figure 17.9) (Data from Ref. 42).
muscles during increases in minute ventilation produced by the rebreathing of CO2, but parasternal intercostal activity remained absent in the males. The number of subjects studied was insufficient to conclude that gender was causal, although the pattern of results in the male subjects is consistent with the results of earlier studies [42]. The mechanisms responsible for these apparent gender differences in the response of extra-diaphragmatic respiratory muscles to anesthesia are unknown. Progesterone is known to affect ventilatory control [116], but there is little information regarding gender-related differences in specific patterns of respiratory muscle use. The original study documenting consistent phasic expiratory muscle activity during nitrous oxide or halothane anesthesia was performed in males [104]. A subsequent study of subjects anesthetized with thiopental and nitrous oxide that included females noted that expiratory activity did not develop in all subjects, but the presence of such activity did not seem to depend on gender [103]. Activity persisted when halothane was added to the inspired gas. Other studies of parasternal intercostal or scalene muscle activity during anesthesia are limited. Tusiewicz et al. [115] found no parasternal intercostal muscle activity in three subjects (gender unspecified) anesthetized with halothane
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during quiet breathing. Drummond [39] found that enflurane administered for a brief period shortly after thiopental induction abolished parasternal and scalene muscle activity in five subjects (gender unspecified). Based on current knowledge, we can only conclude that there is significant variability in the pattern of respiratory muscle activation among anesthetized subjects. Differences among anesthetic agents have not been systematically studied. However, it appears that when added to potent volatile agents, nitrous oxide has little effect on the pattern of activation in human subjects [117]. Diaphragm
The activation of the diaphragm changes during anesthesia, often in unexpected ways. Warner et al. [106] examined the effect of halothane anesthesia on both quiet breathing and on the response of respiratory muscles to CO2 rebreathing. With inspiratory drive to the phrenic motoneurons defined as the rate of rise of EMG activity (EDIA) over the course of inspiration (the closest approximation possible in humans), then drive during quiet breathing is actually increased by halothane anesthesia, accompanied by significant changes in ventilatory timing such as a decrease in the period of breathing. Furthermore, during CO2 rebreathing, the increase in average diaphragm EMG activity for a given increase in PCO2 and the average diaphragm activity at a given PCO2 are both greater during halothane anesthesia compared with wakefulness, suggesting that the response of phrenic motoneuron drive to rebreathing is actually enhanced by halothane anesthesia (Figure 17.11). There are several possible explanations for these somewhat surprising results. During quiet breathing, the duration of diaphragmatic activity is markedly shortened by halothane, and phrenic motoneuron drive (as defined by EDIA) increases so that average EMG activity during quiet breathing is not consistently affected by halothane. Thus, increases in phrenic motoneuron drive could be related to the profound changes in timing produced by halothane. Changes in ventilatory timing during CO2 rebreathing are also affected by anesthesia. For example, breathing frequency does not change in response to rebreathing while anesthetized, so that greater increases in VT are required to produce a given increase in minute ventilation while anesthetized. The need for greater increases in VT may require greater increases in neural drive to the diaphragm. Increased drive to the phrenic nerve during halothane anesthesia may also represent compensation for changes in the activity of other respiratory muscles that normally assist respiration. It is probably overly simplistic to think of the increase in diaphragm activity as a response to the loss of activity in any single muscle group produced by halothane anesthesia. However, the profound effect of halothane anesthesia on
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Figure 17.11 Moving time average of the diaphragm electromyogram (DIA MTA) in one human subject while both awake and anesthetized with 1.2 MAC halothane during CO2 rebreathing. This parameter is a measure of diaphragm activation over the course of inspiration. Values are expressed as a percent of the maximum value during rebreathing while awake and are plotted as a function of expired CO2 tension (left panel) or tidal volume (right panel). Solid lines are linear regressions. Note that the slope of the response as a function of PCO2 is actually increased during anesthesia, and that a greater MTA is required to generate a given tidal volume during anesthesia. This increase in diaphragm activation may represent a response to the respiratory muscle incoordination produced by anesthesia (Data from Ref. 106).
extra-diaphragmatic respiratory muscles raises the possibility that the increase in drive to phrenic motoneurons may represent compensation for the loss of normal respiratory muscle coordination. In the study of Warner et al. [106], analysis of the response of average inspiratory flow (VT/TI), a parameter of overall ventilatory drive, suggests that this compensation was remarkably complete. Although VT/TI during quiet breathing is significantly lower during anesthesia, the slope of the linear regression of inspiratory flow versus minute ventilation was significantly greater while anesthetized (2.8 0.4, mean S.D.) than while awake (1.8 0.1, mean S.D.). Thus, rebreathing produced a greater increase in inspiratory flow for a given change in minute ventilation during anesthesia, despite changes in the activation of extradiaphragmatic muscles. Anesthesia may also alter other aspects of phrenic motoneuron control. While awake, activity in the diaphragm and other inspiratory agonists persists into early expiration. The physiologic significance of this post-inspiratory inspiratory activity (PIIA) is not fully understood, but presumably serves to regulate early expiratory flow and increase mean lung volume [118]. This activity is abolished by halothane anesthesia [106]
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(Figure 17.9). Because PIIA may play a role in producing the normal respiratory rhythm, halothane-induced suppression of PIIA may be related to the halothane-induced changes in ventilatory timing. Summary of anesthetic effects on respiratory muscle activation
In summary, as reviewed in chapters 15 and 16, anesthetic drugs depress global measures of respiratory control such as the resting arterial carbon dioxide tension and the slope of the ventilatory response to carbon dioxide. Sufficient doses of any anesthetic drug will eventually produce profound respiratory depression and apnea. However, consideration of anesthetic effects only in these terms obscures much of the complexity of the drugs’ effects. For example, anesthetic drugs may under some circumstances actually enhance the rebreathing response of neural drive to the primary respiratory muscle, the diaphragm. Thus, anesthetic-induced respiratory depression may be caused by alterations in the distribution and timing of neural drive to the respiratory muscles, rather than a global depression of respiratory motoneuron drive. This respiratory muscle incoordination may have profound consequences for the function of the respiratory pump, as discussed in the following section. Chest Wall Motion Quiet breathing
Clinically, paradoxic inspiratory inward motion of the rib cage was used for many years as a indication that a surgical plane of ether anesthesia had been established [2,119]. In contemporary practice, many clinicians have also noted this pattern of chest wall motion during administration of modern volatile anesthetics. Indeed, initial studies showed that halothane anesthesia reduces the rib cage contribution to tidal volume as measured by changes in thoracic dimensions [94,115]. This was attributed to a selective inhibition of inspiratory intercostal muscle activity by halothane, which normally helps to maintain rib cage expansion, with a relative maintenance of diaphragmatic activity. However, more recent work finds that expansion of the rib cage is actually relatively well preserved during halothane anesthesia with quiet breathing, with only a tendency towards a decrease in its relative contribution to the total change in intrathoracic volume [12]. Other studies of patients anesthetized with isoflurane [120], ketamine [88], or methohexital [87] have also noted a relative preservation of rib cage expansion during quiet breathing. These findings are curious, given the clinical experience above, but recent work provides insight that may explain these seemingly contradictory findings. If inspiration during anesthesia is accomplished primarily by diaphragmatic contraction, we can predict the resultant pattern of rib cage motion. Isolated contraction of the diaphragm has both inspiratory
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and expiratory actions on the rib cage [121]. Inspiratory forces result from the direct action of the diaphragm via its insertions on the rib cage to elevate the ribs and from increases in abdominal pressure acting on the lower rib cage via the area of apposition between diaphragm and the rib cage. Decreases in intrathoracic pressure produced by diaphragm contraction constrict the rib cage. Studies in animals [22,23] and in human subjects [24,26,121] suggest that isolated diaphragmatic activity distorts the rib cage, producing an inward motion of the upper rib cage and an outward motion of the lower rib cage during inspiration. However, this pattern has not been observed during halothane anesthesia [12], suggesting that other muscles act to expand the rib cage during anesthesia. There are two possibilities. First, continued activation of respiratory muscles with inspiratory actions (such as the activity in scalene muscles observed in some subjects) may help maintain rib cage expansion. For example, studies in quadriplegics, and in normal subjects with paralysis of intrinsic rib cage muscles produced by epidural anesthesia, show that continued action of accessory muscles such as the scalenes is sufficient to maintain rib cage expansion during quiet breathing [26,121,122]. Second, cessation of activity in intrinsic muscles with expiratory actions such as the internal intercostal muscles will cause passive rib cage expansion. This activity (when present) is mechanically significant during halothane anesthesia [42], contributing significantly to rib cage expansion (Figure 17.12). The relative importance of these two mechanisms depends on the particular pattern of respiratory activation during anesthesia present in individual patients. Because of the difficulties in measuring motion of the diaphragm in vivo, there is relatively little information regarding its behavior during anesthesia. However, it appears that at least during quiet breathing, the overall pattern of diaphragm motion is maintained during anesthesia, with relatively more axial motion in dependent regions of the diaphragm in subjects lying supine [11–13]. Loaded breathing
Although inspiratory rib cage expansion is relatively well preserved in anesthetized subjects breathing quietly, it may be vulnerable to respiratory loads. The most well studied loading condition is that presented by the rebreathing of carbon dioxide. By measuring the relative contribution of the rib cage and diaphragm–abdomen to tidal volume during CO2 rebreathing, the response of both compartments can be quantified. The compartmental response of the rib cage, quantified as the slope of the relationship between compartmental minute ventilation and PCO2, is significantly reduced by halothane anesthesia [106,115]. This is accompanied by marked asynchrony between rib cage and abdominal motion (which is normally well-coordinated during rebreathing in awake subjects). For
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Figure 17.12 EMGs, gastric and esophageal pressures, and chest wall motion (measured with rib cage and abdominal bands, see Figure 17.3) in a male subject anesthetized with 1.2 MAC end-tidal halothane at the onset (left panel) and conclusion (middle panel) of CO2 rebreathing, which produces hyperpnea. Open and closed circles denote the beginning and end of inspiratory gas flow, respectively. Note recruitment of all respiratory muscles, including the development of phasic expiratory activity in the diaphragm electrode that represents expiratory activity of the adjacent internal intercostal muscle. At the conclusion of rebreathing, paradoxical rib cage motion is present, with rib cage expansion at the onset of expiration. Also shown (right panel) is a unique pattern of breathing that developed in this subject during recovery from rebreathing, in which expiratory muscle activity was absent. Note that in the absence of such activity, there is now paradoxic motion of the rib cage throughout inspiration, with the rib cage decreasing in area as inspiration proceeds. This finding demonstrates the importance of expiratory muscle activity in maintaining rib cage expansion during halothane anesthesia, when inspiratory intercostal muscle activity is suppressed (Data from Ref. 106).
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example, the rib cage moves outward in early expiration during rebreathing while anesthetized, probably because intrathoracic pressure rapidly increases in early expiration as diaphragmatic contraction ends [106] (see Figure 17.12). The lack of PIIA activity in inspiratory muscles during halothane anesthesia accentuates this sudden increase in intrathoracic pressure during early expiration and thus also contributes to this paradoxical rib cage motion. This lack of normal coordination among respiratory muscles contributes to halothane’s ability to depress the overall minute ventilatory response to CO2. For example, when respiratory drive to the diaphragm (defined by the rate of rise of EMG activity) is analyzed as a function of tidal volume, significantly greater EMG activity is needed to generate a given VT during anesthetized rebreathing compared with rebreathing while awake [106] (Figure 17.11). This finding suggests a decrease in the efficiency of diaphragmatic contraction during halothane anesthesia; i.e., a decrease in the volume of gas displaced for a given neural input to the diaphragm. Clinically, the most common load to breathing is presented by increases in upper airway resistance accompanying anesthesia, which may be at least partially responsible for the paradoxic rib cage retraction noted during clinical anesthesia. However, Moote et al. [123] found that resistive loading produced immediate decreases in rib cage expansion in halothaneanesthetized humans only when relatively large loads were imposed, with wide variability in the response of individual subjects. Thus, as in the case of CO2 rebreathing, chest wall responses are relatively well preserved, even in the presence of significant changes in the pattern of respiratory muscle activation. Summary of anesthetic effects on chest wall motion
For the potent volatile agents, the best-studied class of general anesthetics, the pattern of chest wall motion is remarkably well-preserved with the induction of anesthesia, despite significant alterations in the pattern of respiratory muscle activation. Some changes in activation, such as increases in the rate of diaphragm EMG activity, may occur in response to the respiratory muscle incoordination that develops during anesthesia. Significant alterations in motion are seen under conditions of loaded breathing, but the respiratory system is still able to mount a relatively effective response at surgical levels of anesthesia.
IV.
Effects of Anesthesia on Upper Airway Mechanics
In addition to effects on the chest wall musculature, anesthetic drugs and adjuvants such as neuromuscular blocking agents also have important effects on the muscles of the upper airway, which have profound
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consequences for the clinical conduct of anesthesia. Indeed, a large portion of anesthetic practice is devoted to maintaining upper airway patency. A. Activation of Upper Airway Muscles
The depth and choice of anesthetic are critical determinants of activity in the upper airway muscles in experimental animal preparations, and in fact represent a considerable challenge to investigators studying the physiology of upper airway muscles [124]. In general, anesthetic drugs depress such activity. Laryngeal motoneurons with expiratory activities are depressed by pentobarbital [125,126]. Upper airway muscles with phasic inspiratory activity are also depressed by barbiturates [127–131], although one investigator found increases in activity when pentobarbital was given to decerebrate cats [126]. Several reports have documented differential sensitivity to anesthetic depression between chest wall and upper airway muscle motoneuron activities (Figure 17.13) [129–131]. Halothane, enflurane, diazepam, and thiopental all produce a greater depression of hypoglossal nerve activity compared with phrenic nerve activity in paralyzed, ventilated, vagotomized cats [129,130]. Measurements of EMG activities in intact anesthetized cats breathing spontaneously show similar results; EMG activity in the diaphragm is more resistant to depression by halothane than the genioglossus muscle [131]. This differential suppression may be less pronounced after ketamine administration [128,130,132], suggesting that it is not a property common to all general anesthetics. Other reports also have noted apparent preservation of upper airway motoneuron activities during ketamine anesthesia [127,133]. There may be differential sensitivity to anesthesia among individual upper airway muscles [129,134]. For example, Nishino et al. [129] found that halothane and enflurane depressed hypoglossal nerve activity more than recurrent laryngeal nerve activity. Studies of anesthetic effects on the activation of upper airway muscles in human subjects pose particular challenges for at least two reasons. First, it is often technically difficult to obtain EMG activities in human subjects (the genioglossus muscle is most often used as a representative pharyngeal dilator). Second, many factors influence the activity of upper airway muscles, including blood gases, state of arousal, age, blood pressure, temperature, lung volume, and reflexes mediated by pharyngeal pressure [135–142]. Thus, it may be difficult to distinguish between primary effects of anesthetic drugs (in the absence of other changes), and secondary effects of the drugs (e.g., changes in FRC). Some studies have found a differential effect of anesthesia on the temporal pattern of activity in upper airway muscles that dilate the pharynx in human subjects. In general, tonic activity is diminished by all anesthetics examined, whereas phasic activity may be relatively well maintained with
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Figure 17.13 Phasic inspiratory muscle electromyogram activity (expressed as a percent of peak moving time average) as a function of halothane concentration for the diaphragm (DI), intercostal (IC), and the genioglossus (GG) muscles in spontaneously breathing cats. Note that the activity of the upper airway muscle was preferentially suppressed as anesthetic depth was increased (redrawn from Ref. 131).
some anesthetic agents. Drummond [143] found that incremental doses of intravenous thiopental decreased the predominantly tonic activity present while awake in the genioglossus and strap muscles of the neck (measured with surface electrodes), with the appearance of phasic activity in many subjects. Similar results have been noted with midazolam [133,144]. These studies noted that phasic activity was often able to increase in response to partial airway obstruction. The potent volatile anesthetics may have more profound depressing effects on upper airway muscle activity, although direct comparisons have not been performed and data are scarce. Eastwood et al.
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[145] found that low doses of isoflurane (0.4% end-tidal) almost completely eliminated both tonic and phasic genioglossus EMG activity (measured by intramuscular electrodes) during spontaneous breathing. In addition, there was no activation of activity in response to decreases in mouth pressure (sufficient to cause upper airway collapse). Such decreases cause significant reflex activation of upper airway muscles during wakefulness [146] and natural sleep [147]. These results show that isoflurane causes profound depression of upper airway muscle control. This finding is consistent with animal studies showing inhibition of laryngeal receptors that mediate upper airway reflexes by volatile anesthetics, which differ in potency for this effect [147,148]. B. Upper Airway Mechanics
How changes in the activity of upper airway muscles influence upper airway caliber is complex and poorly understood. Basic descriptive information both of morphology (provided by imaging techniques and fiberoptic endoscopy) and of function (such as upper airway pressure-flow relationships [150]) during anesthesia is only now being generated. Although anesthetic effects on upper airway function may not mimic those of natural sleep, several important principles observed during natural sleep may also apply during anesthesia. Because the upper airway is a collapsible tube that is subjected to subatmospheric pressure during inspiration, upper airway muscles presumably stiffen the airway to resist collapse [47,54,150–152]. However, there is no consistent relationship between phasic EMG activity of upper airway muscles and upper airway resistance. For example, resistance changes little from non-REM to REM sleep [153], even though upper airway EMG activity is reduced [154]. Similarly, Drummond [39] found that no reduction in the activity of upper airway muscles could be related directly to the onset of airway obstruction; rather, activity was often increased, presumably in an attempt to overcome partial obstruction. Thus airway obstruction may not be caused by just a simple diminution of activity but by disruption of the normal coordination of activity of muscles controlling different segments of the airways [39,155]. Disruption of the normal coordination between upper airway and chest wall muscle activities may also contribute to sleep-disordered breathing [156]; this possibility has not been investigated for the anesthetic drugs. Regardless of mechanism, several common themes have emerged in recent studies of the effects of anesthetics on upper airway mechanics. First, although the luminal dimensions of the pharynx are decreased at several levels, the site that is the most narrowed during anesthesia is not at the base of the tongue [157], as is often supposed, but at the level of the soft palate (Figure 17.14) [157–162]. The mechanism by which this occurs is not known, but it can also lead to collapse of more distal pharyngeal areas as inspiration
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Figure 17.14 Diagrammatic representation of mid-sagittal section through the upper airway, showing changes in upper airway anatomy during apnea following the induction of thiopental anesthesia. Note that although all dimensions are lessened, the primary site of occlusion is at the soft palate (redrawn from Ref. 158).
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produces negative inspiratory pressures. The site of obstruction may also differ among types of patients. For example, patients with obstructive sleep apnea may be more likely than normal patients to develop additional obstruction at the tongue level with propofol sedation [163]. Second, anesthesia and sedation enhance the collapsibility of the upper airway, defined as the ability of negative pharyngeal pressures to produce airway occlusion [144,164]. This is certainly related in part to decreases in activation of pharyngeal dilators by anesthetics, as loss of muscle tone produced by profound neuromuscular blockade (but not mild blockade) increases collapsibility [165,166]. However, this is not the whole story. Eastwood et al. [145] found that upper airway collapsibility increased with increasing depth of isoflurane anesthesia, despite the fact that little genioglossus EMG activity was noted under any condition while subjects were anesthetized with as little as 0.4% end-tidal isoflurane (Figure 17.15). From concurrent measurements of esophageal pressures, they inferred that lung volume decreased as anesthetic depth increased, and that this factor decreased longitudinal airway wall tension, increasing collapsibility. Thus, effects of anesthesia on the FRC may have secondary effects on the behavior of the upper airway. Finally, the role of tonic vs. phasic activity in the maintenance of airway patency during anesthesia remains to be defined; current evidence suggests that loss of tonic activity may be especially important, as continued phasic activity during anesthesia and sedation does not prevent the occurrence of obstruction [39,144]. Recent studies have provided the structural basis for maneuvers used to maintain upper airway patency in clinical practice [158,159,161,168]. For example, forward advancement of the mandible increases the area of the glottic opening; continuous positive airway pressure has similar effects [170]. C. Summary of Anesthetic Effects on Upper Airway Mechanics
Anesthetic drugs have profound effects on the activation of upper airway muscles, even when administered in low doses. These changes in control produce an increase in the pharyngeal collapsibility, although the exact mechanisms by which this occurs remain obscure. V.
Effects of Anesthesia on Lung Mechanics
Some anesthetic drugs may have primary effects on several elements of the lung that affect its mechanical properties, such as airway smooth muscle and the surfactant lining the airways. However, many aspects of the mechanical behavior of the lung, such as the regional distribution of ventilation, are in large measure determined by its container, the chest
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Figure 17.15 Airway collapsibility in human subjects as a function of expired isoflurane concentration during mask anesthesia with spontaneous breathing. Airway pressure was progressively reduced from a level at which no flow limitation occurred (11.8 2.7 cm H2O), and the pressure-flow relationship extrapolated to zero flow to calculate the airway pressure at which complete collapse occurred (critical airway pressure). Note that critical airway pressure increased with anesthetic depth, despite no significant change in tonic genioglossus EMG activity (which was minimal throughout the study compared with activity while awake). Values are mean S.D.; n ¼ 16; * denotes significant difference from value at 0.4% end-tidal isoflurane (Data from Ref. 145.)
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wall. It appears that many of the changes in lung mechanics caused by anesthesia are secondary to changes in chest wall mechanics, rather than primary effects on the lung itself. This section will concentrate on these secondary changes. A. Atelectasis and Gas Exchange
An important series of investigations from Hedenstierna and colleagues [110,170–185] found that anesthesia consistently produces densities in the dependent regions of the lung as imaged by computed tomography (Figure 17.16). The extent of these densities, thought to represent areas of atelectasis, is correlated with the magnitude of shunt produced by anesthesia [171,177,179,186,187]. The densities are observed within minutes of anesthetic induction of patients with both volatile and intravenous anesthetic agents (with the exception of ketamine [188]) during both spontaneous breathing and mechanical ventilation. Atelectasis can be lessened with positive end-expiratory pressure [171,173,175,176,184], and can be largely reversed by vigorous expansile maneuvers [179,188,189] (e.g., positive airway pressures of 40 cm H2O held for 15 sec). The extent of atelectasis is dependent on the nature of inspired gas, with high concentrations of oxygen promoting its formation [180–182,190,191]. These atelectatic areas may represent a significant source of postoperative pulmonary morbidity in some patients, as they may persist for some time after surgery [172,178,190,192]. Atelectasis is almost certainly caused by anesthetic-induced changes in chest wall mechanics, yet the mechanism by which this occurs is not known. It has been hypothesized that preferential cephalad displacement of the diaphragm compresses lung tissue in dependent regions [175]. Contraction of the diaphragm produced by phrenic nerve stimulation in anesthetized subjects reduces lung atelectasis [110], suggesting that changes in chest wall shape produced by alterations in diaphragm muscle tone can indeed affect the extent of atelectasis. If atelectasis is caused by compression of lung parenchyma by a cephalad displacement of the dependent diaphragm, then the greater the displacement, the greater should be the amount of atelectasis. However, two studies have found no such correlation (Figure 17.17) [110,193]. Thus, although there is little doubt that atelectasis is caused by anesthesia-induced changes in chest wall shape, the cause is not as simple as a cephalad shift of the diaphragm. This lack of correlation between the amount of atelectasis and anesthesia-induced changes in endexpiratory position of any chest wall structure implies that the formation of atelectasis cannot be ascribed to any single chest wall element in these healthy subjects. In addition to atelectasis, changes in regional ventilation caused by anesthesia have also been attributed specifically to changes in diaphragm
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Figure 17.16 Two-dimensional representation of a volume image of the thorax, obtained using fast volumetric CT scanning, of a human subject anesthetized with 1.2 MAC halothane. The surface of the lung is shown in shades of gray and areas of atelectasis are shown in white. Conceptually, this represents a view of a gray transparent 3-dimensional model of lung shape, with a superimposed opaque white model of atelectasis. The anteroposterior view (top) clearly shows the right and left lungs, with a visible cardiac shadow. The lateral view (bottom) demonstrates that the atelectatic areas are located in the most dependent lung regions in this subject lying supine. In this view, down is posterior, and left is caudad (Data from Ref. 194).
motion. In the supine position, the institution of anesthesia with paralysis and mechanical ventilation increases regional ventilation in nondependent lung regions relative to dependent lung regions (Figure 17.18) [195]. This change in the distribution of ventilation has been explained simply by an increase in the axial motion of nondependent regions of the diaphragm. However, when in the prone position, ventilation both while awake and
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Atelectasis, % total lung volume
5 Spontaneous breathing, rs = 0.16 4
Mechanical ventilation, rs = 0.69
3 2 1 0 0.0
0.5
1.0
1.5
2.0
2.5
3.0
Change in average dependent diaphragm position, cm
Figure 17.17 Relationship between the volume of atelectasis, measured from images such as Figure 17.16 and expressed as a percentage of total lung volume, and the change in the average axial position of the dependent regions of the diaphragm at end-expiration with the induction of 1.2 MAC halothane anesthesia in human subjects (see Figure 17.7 for method used to calculate). Positive values denote that the average position moved cephalad (headward). Values measured during both spontaneous breathing and mechanical ventilation are shown along with the Spearman correlation coefficients (rs) for these relationships. Note that there is no significant correlation between displacement of the dependent diaphragm and atelectasis, suggesting that atelectasis is not formed simply by such displacement (Data from Ref. 194).
anesthetized is still preferentially distributed to dependent regions, whereas most axial motion of the diaphragm is in nondependent regions [196]. Thus, regional ventilation is not determined only by the motion of the adjacent diaphragm. These two findings should not be surprising, considering the complexity of the factors that determine the stresses applied to the lung tissue, and hence to regional lung expansion [197]. These factors include the weight of the lung tissue, the interaction between the lung and its container (which have intrinsically different shapes), and the mechanical effects of imbedded bronchi and blood vessels. There is good evidence that pronounced distortion of the chest wall changes regional lung expansion in humans [198,199]. However, less pronounced changes in chest wall shape in supine humans might have little effect on regional lung expansion [200]. Although anesthesia-induced changes in the shape of the chest wall may change the distribution of stresses applied
General Anesthesia and Respiratory Mechanics
719 Prone
Supine Top Vertical distance down the lung, cm
0
Awake Anesth-paral
10
20 Bottom 50
100
150
50
100
150
Ventilation index, %
Cephalad
Spine
Figure 17.18 Regional ventilation in human subjects as a function of vertical distance from the most anterior lung regions while awake (breathing spontaneously) and anesthetized-paralyzed (mechanically ventilated) in the supine (left panel) and prone (right panel) positions. Also shown (lower panel) are the average axial displacements of the diaphragm during tidal breathing in both conditions, measured by fast volumetric CT scanning. In these diagrams, the solid lines indicate the endexpiratory and end-inspiratory positions of the diaphragm while awake, and the stippled areas represent the excursion of the diaphragm produced by mechanical ventilation while anesthetized-paralyzed. Note that in the prone position, ventilation is still preferentially distributed to the dependent regions, whereas most of the displacement of the diaphragm occurs in non-dependent regions. This demonstrates that regional ventilation is not determined only by the motion of the adjacent diaphragm (Data derived from Refs. 9 and 196).
to the lung, changes in regional lung expansion cannot be simply predicted by the local motion of the adjacent chest wall [201]. At present, it can only be concluded that the interaction of several potentially significant factors, including the shape of chest wall structures (such as the thoracic spine, rib cage, diaphragm) and the volume or distribution of blood in the thorax, leads to atelectasis and changes in regional ventilation.
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Although atelectasis is an important factor in anesthesia-induced changes in pulmonary mechanics, two caveats apply. First, not all of the decrease in FRC produced by anesthesia can be accounted for by atelectasis [184]. Using volumetric measurements obtained with 3-dimensional CT scanning of anesthetized human subjects, Warner et al. [194] estimated that only about 25% of the decrease in FRC produced by halothane anesthesia could be attributed to atelectatic areas, with the remainder of the FRC decrease caused by changes in intrathoracic blood volume and decreases in volume of lung regions that remained aerated. Second, atelectasis is not the sole cause of gas-exchange abnormalities during and after anesthesia, and other changes in regional ventilation associated with changes in chest wall mechanics, such as those described above (Figure 17.18), may be important [185,192,194,195]. B. Elastic Behavior
Anesthetic effects on the lung mechanics can be quantified by measuring the relationship between pressures applied to the respiratory system and consequent changes in lung volume. In general, the elastic recoil of the total respiratory system increases with the induction of anesthesia [82,202], so that a greater inflation pressure is required to maintain a given lung volume. This increase is affected neither by the depth of anesthesia nor by the addition of paralysis to anesthesia, and is not progressive with time. These changes in recoil are associated with decreases in both the static and dynamic compliances of the respiratory system. The primary source of this decreased compliance is the lung (Figure 17.19) [82]. Chest wall compliance appears to be little affected over most of the range of lung volumes; at low lung volumes chest wall recoil may be decreased, although it is technically difficult to estimate pleural pressures at low lung volumes in subjects lying supine [82]. Anesthesia could have a primary action on the lung by stimulating smooth muscle or other contractile elements in the airways or lung parenchyma, by directly causing atelectasis, by changing surfactant composition [203–205], or by several other possibilities. However, studies to date do not strongly support the importance of any of these mechanisms in vivo. Rather, alterations in lung compliance are most likely secondary to changes in chest wall function, especially as lung elastic properties are quite dependent on the conditions of lung ventilation. One example of these changes is the atelectasis discussed above. In addition, conditions where ventilation occurs at low lung volumes, such as produced by external strapping of the chest wall, are associated with a progressive decrease in lung compliance [206], even in the absence of atelectasis. Changes in surfactant properties have been advanced to explain this decrease [207]. Thus, the primary effect of anesthesia is to change the shape and motion
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Pentothal, Meperidine
Volume, % TLC control
Total system
Lung
Chest wall
100 80 60 40 20 0
0 10 20 30 Pao, cm H2O Awake
0
Anesth.
10 20 30 PL, cm H2O
0 10 20 Pes, cm H2O
Anesth. paral.
Figure 17.19 Mean pressure-volume relationships measured in five human subjects lying supine while awake, anesthetized with thiopental and meperidine, and anesthetized with paralysis. Separate lung and chest wall compliances were determined using measurements of esophageal pressure as an estimate of average intrathoracic pressure. Note that anesthesia, with or without paralysis, decreased total respiratory system compliance, an effect primarily attributable to decreases in lung compliance (Data from Ref. 82).
of the chest wall, leading to secondary changes in lung elastic behavior [77,82]. C. Pressure-flow Relationships
In contrast to elastic behavior, effects of anesthesia on pressure-flow relationships in the lung (as quantified by parameters such as airway resistance) are primarily caused by effects on intrinsic elements of the lung, such as airway smooth muscle. Most anesthetics produce bronchodilation via a variety of mechanisms, a topic beyond the scope of this chapter (see [1] for review). Changes in elastic properties also contribute to changes in some measures of pulmonary resistance [208]. Although these primary effects are of principal importance, secondary changes in pressure-flow relationships caused by anesthetic effects on the chest wall are also possible. Normally, airway resistance increases as lung volume decreases, a consequence of the mechanical interaction between the airways and the surrounding lung parenchyma, which tethers the airway with a force proportional to lung volume. However, these interactions become less important as airway smooth muscle tone is reduced. Indeed, the relaxation of smooth muscle produced by many anesthetics minimizes the dependence of airway resistance on lung volume. Thus, such secondary changes are likely to be of little significance [209].
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Summary
Although breathing is often considered to be a relatively simple activity involving primarily a single muscle (the diaphragm), normal breathing actually requires the complex, coordinated action of multiple components of the chest wall and upper airway. This coordination is exquisitely sensitive to disruption by general anesthetics. Although sufficient doses of general anesthetic drugs produce apnea, anesthetic-induced respiratory depression at lower doses (consistent with surgical levels of anesthesia) may be caused by alterations in the distribution and timing of neural drive to the respiratory muscles, rather than a global depression of respiratory motoneuron drive. This loss of the normal coordination among the respiratory muscles in turn affects the expansion of the underlying lung and its ability to exchange gas.
References 1. 2. 3.
4.
5.
6. 7.
8. 9.
10.
Warner, D.O., Airway pharmacology, in Airway Management: Principles and Practice, Benumof, J., ed., St. Louis, J. Mosby Yearbook, pp. 74–101, 1995. Snow, J., On Chloroform and Other Anesthetics: Their Action and Administration, London, John Churchill, 1858. Fishman, A.P., Macklem, P.T. and Mead, J., eds., The respiratory system, mechanics of breathing, in Handbook of Physiology, Sec. 3, Vol. 3, Bethesda, MD, American Physiologic Society, 1986. Warner, D.O., Respiratory muscle function, in Anesthesia: Biological Foundations, Yaksh T.L., Lynch C., III, Zapol W.M., Maze M., Biebuyck J.F., Saidman L.J., eds., New York, Raven Press, pp. 1395–1415, 1997. De Troyer, A. and Loring, S., Action of the respiratory muscles, in Handbook of Physiology, The Respiratory System, Mechanics and Breathing, Fishman A.P., Macklem P.T., Mead S., eds., Bethesda, M.D., American Physiological Society, pp. 443–461, 1986. De Troyer, A., Sampson, M., Sigrist, S. and Macklem, P.T., The diaphragm: two muscles, Science 213, 237–238, 1981. De Troyer, A., Sampson, M., Sigrist, S. and Macklem, P.T., Action of costal and crural parts of the diaphragm on the rib cage in dog, J. Appl. Physiol. 53, 30–39, 1982. Mead, J., Functional significance of the area of apposition of diaphragm to rib cage [proceedings], Am. Rev. Respir. Dis. 119, 31–32, 1979. Krayer, S., Rehder, K., Vettermann, J., Didier, E.P. and Ritman, E.L., Position and motion of the human diaphragm during anesthesia-paralysis, Anesthesiology 70, 891–898, 1989. Whitelaw, W.A., Shape and size of the human diaphragm in vivo, J. Appl. Physiol. 62, 180–186, 1987.
General Anesthesia and Respiratory Mechanics 11. 12.
13.
14.
15. 16. 17.
18.
19. 20.
21. 22.
23.
24.
25.
26.
723
Froese, A.B. and Bryan, A.C., Effects of anesthesia and paralysis on diaphragmatic mechanics in man, Anesthesiology 41, 242–255, 1974. Warner, D.O., Warner, M.A. and Ritman, E.L., Human chest wall function while awake and during halothane anesthesia, I. Quiet breathing, Anesthesiology 82, 6–19, 1995. Kleinman, B.S., Frey, K., VanDrunen, M., Sheikh, T., DiPinto, D., Mason, R. and Smith, T., Motion of the diaphragm in patients with chronic obstructive pulmonary disease while spontaneously breathing versus during positive pressure breathing after anesthesia and neuromuscular blockade, Anesthesiology 97, 298–305, 2002. D’Angelo, E. and Sant’Ambrogio, G., Direct action of contracting diaphragm on the rib cage in rabbits and dogs, J. Appl. Physiol. 36, 715–719, 1974. Loring, S.H. and Mead, J., Action of the diaphragm on the rib cage inferred from a force-balance analysis, J. Appl. Physiol. 53, 756–760, 1982. Mortola, J.P. and Sant’Ambrogio, G., Motion of the rib cage and the abdomen in tetraplegic patients, Clin. Sci. Mol. Med. 54, 25–32, 1978. Campbell, E.J.M. and Newsom-Davis, J., The intercostal muscles and other muscles of the rib cage, in The Respiratory Muscles: Mechanics and Neural Control, Campbell, E.J.M., Agostoni, E. and Newsom-Davis, J., eds., Philadelphia, Saunders, pp. 161–174, 1970. De Troyer, A., Kelly, S., Macklem, P.T. and Zin, W.A., Mechanics of intercostal space and actions of external and internal intercostal muscles, J. Clin. Invest. 75, 850–857, 1985. DiMarco, A.F., Romaniuk, J.R. and Supinski, G.S., Action of the intercostal muscles on the rib cage, Respir. Physiol. 82, 295–306, 1990. DiMarco, A.F., Romaniuk, J.R. and Supinski, G.S., Mechanical action of the interosseous intercostal muscles as a function of lung volume, Am. Rev. Respir. Dis. 142, 1041–1046, 1990. Wilson, T.A. and De Troyer, A., Effect of respiratory muscle tension on lung volume, J. Appl. Physiol. 73, 2283–2288, 1992. Krayer, S., Decramer, M., Vettermann, J., Ritman, E.L. and Rehder, K., Volume quantification of chest wall motion in dogs, J. Appl. Physiol. 65, 2213–2220, 1988. Warner, D.O., Brichant, J.F., Ritman, E.L. and Rehder, K., Chest wall motion during epidural anesthesia in dogs, J. Appl. Physiol. 70, 539–547, 1991. De Troyer, A. and Heilporn, A., Respiratory mechanics in quadriplegia, The respiratory function of the intercostal muscles, Am. Rev. Respir. Dis. 122, 591–600, 1980. Gandevia, S.C., Leeper, J.B., McKenzie, D.K. and De Troyer, A., Discharge frequencies of parasternal intercostal and scalene motor units during breathing in normal and COPD subjects, Am. J. Respir. Crit. Care Med. 153, 622–628, 1996. De Troyer, A. and Estenne, M., Coordination between rib cage muscles and diaphragm during quiet breathing in humans, J. Appl. Physiol. 57, 899– 906, 1984.
724 27.
28.
29. 30. 31.
32. 33. 34. 35.
36.
37. 38. 39. 40. 41. 42.
43.
44.
45.
Warner Gilroy, R.J. Jr., Lavietes, M.H., Loring, S.H., Mangura, B.T. and Mead, J., Respiratory mechanical effects of abdominal distension, J. Appl. Physiol. 58, 1997–2003, 1985. Mier, A., Brophy, C., Estenne, M., Moxham, J., Green, M. and De Troyer, A., Action of abdominal muscles on rib cage in humans, J. Appl. Physiol. 58, 1438– 1443, 1985. De Troyer, A., Sampson, M., Sigrist, S. and Kelly, S., How the abdominal muscles act on the rib cage, J. Appl. Physiol. 54, 465–469, 1983. Konno, K. and Mead, J., Measurement of the separate volume changes of rib cage and abdomen during breathing, J. Appl. Physiol. 22, 407–422, 1967. Loring, S.H., Mead, J. and Griscom, N.T., Dependence of diaphragmatic length on lung volume and thoracoabdominal configuration, J. Appl. Physiol. 59, 1961–1970, 1985. Mead, J. and Loring, S.H., Analysis of volume displacement and length changes of the diaphragm during breathing, J. Appl. Physiol. 53, 750–755, 1982. Kim, M.J., Druz, W.S., Danon, J., Machnach, W. and Sharp, J.T., Mechanics of the canine diaphragm, J. Appl. Physiol. 41, 369–382, 1976. Laporta, D. and Grassino, A., Assessment of transdiaphragmatic pressure in humans, J. Appl. Physiol. 58, 1469–1476, 1985. Duggan, J.E. and Drummond, G.B., Abdominal muscle activity and intraabdominal pressure after upper abdominal surgery, Anesth. Analg. 69, 598–603, 1989. Agostoni, E. and Hyatt, R.E., Static behavior of the respiratory system, in Handbook of Physiology, The Respiratory System, Mechanics of Breathing, Sec. 3, Vol. 3. Fishman, A.P., Macklem, P.T. and Mead, J., eds., Bethesda, MD, American Physiological Society, pp. 113–130, 1986. De Troyer, A., Bastenier, J. and Delhez, L., Function of respiratory muscles during partial curarization in humans, J. Appl. Physiol. 49, 1049–1056, 1980. De Troyer, A. and Sampson, M.G., Activation of the parasternal intercostals during breathing efforts in human subjects, J. Appl. Physiol. 52, 524–529, 1982. Drummond, G.B., Reduction of tonic rib cage muscle activity by anesthesia with thiopental, Anesthesiology 67, 695–700, 1987. Druz, W.S. and Sharp, J.T., Activity of respiratory muscles in upright and recumbent humans, J. Appl. Physiol. 51, 1552–1561, 1981. Muller, N., Volgyesi, G., Becker, L., Bryan, M.H. and Bryan, A.C., Diaphragmatic muscle tone, J. Appl. Physiol. 47, 279–284, 1979. Warner, D.O., Warner, M.A. and Ritman, E.L., Mechanical significance of respiratory muscle activity in humans during halothane anesthesia, Anesthesiology 84, 309–321, 1996. Colin, A.A., Wohl, M.E., Mead, J., Ratjen, F.A., Glass, G. and Stark, A.R., Transition from dynamically maintained to relaxed end-expiratory volume in human infants, J. Appl. Physiol. 67, 2107–2111, 1989. Kosch, P.C., Hutchinson, A.A., Wozniak, J.A., Carlo, W.A. and Stark, A.R., Posterior cricoarytenoid and diaphragm activities during tidal breathing in neonates, J. Appl. Physiol. 64, 1968–1978, 1988. Kosch, P.C. and Stark, A.R., Dynamic maintenance of end-expiratory lung volume in full-term infants, J. Appl. Physiol. 57, 1126–1133, 1984.
General Anesthesia and Respiratory Mechanics 46.
47.
48. 49.
50.
51.
52.
53.
54.
55.
56. 57.
58. 59.
60.
61. 62.
725
van Lunteren, E. and Strohl, K.P., Striated respiratory muscles of the upper airways, in Respiratory Function of the Upper Airway, Campbell E.J.M., Agostoni E. and Newsom-Davis J., eds., New York, Marcel Dekker, Inc., 1988, pp. 87–123. Mathew, O.P., Abu-Osba, Y.K. and Thach, B.T., Influence of upper airway pressure changes on genioglossus muscle respiratory activity, J. Appl. Physiol. 52, 438–444, 1982. Andrew, B.L., The respiratory displacement of the larynx: a study of the innervation of accessory respiratory muscles, J. Physiol. 130, 474–487, 1955. Roberts, J.L., Reed, W.R. and Thach, B.T., Pharyngeal airway-stabilizing function of sternohyoid and sternothyroid muscles in the rabbit, J. Appl. Physiol. 57, 1790–1795, 1984. Brancatisano, A., Dodd, D.S. and Engel, L.A., Posterior cricoarytenoid activity and glottic size during hyperpnea in humans, J. Appl. Physiol. 71, 977–982, 1991. Kuna, S.T., Smickley, J.S. and Insalaco, G., Posterior cricoarytenoid muscle activity during wakefulness and sleep in normal adults, J. Appl. Physiol. 68, 1746–1754, 1990. Wheatley, J.R., Brancatisano, A. and Engel, L.A., Respiratory-related activity of cricothyroid muscle in awake normal humans, J. Appl. Physiol. 70, 2226–2232, 1991. Kuna, S.T., Insalaco, G. and Woodson, G.E., Thyroarytenoid muscle activity during wakefulness and sleep in normal adults, J. Appl. Physiol. 65, 1332–1339, 1988. Wheatley, J.R., Kelly, W.T., Tully, A. and Engel, L.A., Pressure-diameter relationships of the upper airway in awake supine subjects, J. Appl. Physiol. 70, 2242–2251, 1991. Hairston, L.E. and Sauerland, E.K., Electromyography of the human palate: discharge patterns of the levator and tensor veli palatini, Electromyogr. Clin. Neurophysiol. 21, 287–297, 1981. Remmers, J.E., deGroot, W.J., Sauerland, E.K. and Anch, A.M., Pathogenesis of upper airway occlusion during sleep, J. Appl. Physiol. 44, 931–938, 1978. Sauerland, E.K. and Harper, R.M., The human tongue during sleep: electromyographic activity of the genioglossus muscle, Exp. Neurol. 51, 160–170, 1976. Brancatisano, T., Collett, P.W. and Engel, L.A., Respiratory movements of the vocal cords, J. Appl. Physiol. 54, 1269–1276, 1983. Brancatisano, T.P., Dodd, D.S. and Engel, L.A., Respiratory activity of posterior cricoarytenoid muscle and vocal cords in humans, J. Appl. Physiol. 57, 1143–1149, 1984. Tully, A., Brancatisano, A., Loring, S.H. and Engel, L.A., Influence of posterior cricoarytenoid muscle activity on pressure-flow relationship of the larynx, J. Appl. Physiol. 70, 2252–2258, 1991. England, S.J. and Bartlett, D., Jr., Changes in respiratory movements of the human vocal cords during hyperpnea, J. Appl. Physiol. 52, 780–785, 1982. Insalaco, G., Kuna, S.T., Cibella, F. and Villeponteaux, R.D., Thyroarytenoid muscle activity during hypoxia, hypercapnia, and voluntary hyperventilation in humans, J. Appl. Physiol. 69, 268–273, 1990.
726 63.
64. 65. 66. 67. 68.
69.
70.
71.
72.
73.
74. 75. 76.
77.
78.
79.
Warner Rehder, K. and Marsh, H.M., Respiratory mechanics during anesthesia and mechanical ventilation, in Handbook of Physiology, The Respiratory System, Mechanics of Breathing. Sec. 3, Vol. 3. Fishman, A.P., Macklem, P.T. and Mead, J., eds., Bethesda, MD, American Physiological Society, pp. 737–752, 1986. De Troyer, A., Gilmartin, J.J. and Ninane, V., Abdominal muscle use during breathing in unanesthetized dogs, J. Appl. Physiol. 66, 20–27, 1989. De Troyer, A. and Ninane, V., Triangularis sterni: a primary muscle of breathing in the dog, J. Appl. Physiol. 60, 14–21, 1986. Gilmartin, J.J, Ninane, V. and De Troyer, A., Abdominal muscle use during breathing in the anesthetized dog, Respir. Physiol. 70, 159–171, 1987. Ninane, V., Baer, R.E. and De Troyer, A. Mechanism of triangularis sterni shortening during expiration in dogs, J. Appl. Physiol. 66, 2287–2292, 1989. Warner, D.O., Joyner, M.J. and Rehder, K., Electrical activation of expiratory muscles increases with time in pentobarbital-anesthetized dogs, J. Appl. Physiol. 72, 2285–2291, 1992. Warner, D.O., Krayer, S., Rehder, K. and Ritman, E.L., Chest wall motion during spontaneous breathing and mechanical ventilation in dogs, J. Appl. Physiol. 66, 1179–1189, 1989. Schroeder, M.A., Tao, H.Y. and Farkas, G.A., Mechanical role of expiratory muscle recruitment during eupnea in supine anesthetized dogs, J. Appl. Physiol. 70, 2025–2031, 1991. Farkas, G.A. and Schroeder, M.A., Functional significance of expiratory muscles during spontaneous breathing in anesthetized dogs, J. Appl. Physiol. 74, 238–244, 1993. Farkas, G.A. and Schroeder, M.A., Mechanical role of expiratory muscles during breathing in prone anesthetized dogs, J. Appl. Physiol. 69, 2137–2142, 1990. Henke, K.G., Sharratt, M., Pegelow, D. and Dempsey, J.A., Regulation of end-expiratory lung volume during exercise, J. Appl. Physiol. 64, 135–146, 1988. Warner, D.O., Joyner, M.J. and Ritman, E.L., Anesthesia and chest wall function in dogs, J. Appl. Physiol. 76, 2802–2813, 1994. Warner, D.O., Joyner, M.J. and Ritman, E.L., Chest wall responses to rebreathing in halothane-anesthetized dogs, Anesthesiology 83, 835–843, 1995. Rich, C.R., Rehder, K., Knopp, T.J. and Hyatt, R.E., Halothane and enflurane anesthesia and respiratory mechanics in prone dogs, J. Appl. Physiol. 46, 646–653, 1979. Warner, D.O. and Rehder, K., Influence of anesthesia on the thorax, in The Thorax, 2nd ed., Roussos, C., ed., New York, Marcel Dekker, Inc., pp. 1585– 1598, 1995. Bergman, N.A., Reduction in resting end-expiratory position of the respiratory system with induction of anesthesia and neuromuscular paralysis, Anesthesiology 57, 14–17, 1982. Don, H.F., Wahba, W.M. and Craig, D.B., Airway closure, gas trapping, and the functional residual capacity during anesthesia, Anesthesiology 36, 533–539, 1972.
General Anesthesia and Respiratory Mechanics 80.
81.
82.
83.
84.
85.
86.
87.
88.
89.
90.
91.
92.
93.
94.
727
Hewlett, A.M., Hulands, G.H., Nunn, J.F. and Milledge, J.S., Functional residual capacity during anaesthesia III: Artificial ventilation, Br. J. Anaesth. 46, 495–503, 1974. Howell, J.B.L. and Peckett, B.W., Studies of the elastic properties of the thorax of supine anaesthetized paralysed human subjects, J. Physiol. 136, 1–19, 1957. Westbrook, P.R., Stubbs, S.E., Sessler, A.D., Rehder, K. and Hyatt, R.E., Effects of anesthesia and muscle paralysis on respiratory mechanics in normal man, J. Appl. Physiol. 34, 81–86, 1973. Rehder, K., Sittipong, R. and Sessler, A.D., The effects of thiopentalmeperidine anesthesia with succinylcholine paralysis on functional residual capacity and dynamic lung compliance in normal sitting man, Anesthesiology 37, 395–398, 1972. Pelosi, P., Croci, M., Ravagnan, I., Cerisara, M., Vicardi, P., Lissoni, A. and Gattiononi, L., Respiratory system mechanics in sedated, paralyzed, morbidly obese patients, J. Appl. Physiol. 82, 811–818, 1997. Damia, G., Mascheroni, D., Croci, M. and Tarenzi, L., Perioperative changes in functional residual capacity in morbidly obese patients, Br. J. Anaesth. 60, 574–578, 1988. Pelosi, P., Croci, M., Ravagnan, I., Tredici, S., Pedoto, A., Lissoni, A. and Gattinoni, L., The effects of body mass on lung volumes, respiratory mechanics, and gas exchange during general anesthesia, Anesth. Analg. 87, 654–660, 1998. Bickler, P.E., Dueck, R. and Prutow, R.J., Effects of barbiturate anesthesia on functional residual capacity and ribcage/diaphragm contributions to ventilation, Anesthesiology 66, 147–152, 1987. Mankikian, B., Cantineau, J.P., Sartene, R., Clergue, F. and Viars, P., Ventilatory pattern and chest wall mechanics during ketamine anesthesia in humans, Anesthesiology 65, 492–499, 1986. Rutherford, J.S., Logan, M.R. and Drummond, G.B., Changes in endexpiratory lung volume on induction of anaesthesia with thiopentone or propofol, Br. J. Anaesth. 73, 579–582, 1994. Hedenstierna, G., Strandberg, A., Brismar, B., Lundquist, H., Svensson, L. and Tokics, L., Functional residual capacity, thoracoabdominal dimensions, and central blood volume during general anesthesia with muscle paralysis and mechanical ventilation, Anesthesiology 62, 247–254, 1985. Krayer, S., Rehder, K., Beck, K.C., Cameron, P.D., Didier, E.P. and Hoffman, E.A., Quantification of thoracic volumes by three-dimensional imaging, J. Appl. Physiol. 62, 591–598, 1987. Drummond, G.B., Pye, D.W., Annan, F.J. and Tothill, P., Changes in blood volume distribution between legs and trunk during halothane anaesthesia, Br. J. Anaesth. 75, 707–712, 1995. Hedenstierna, G., Lofstrom, B. and Lundh, R., Thoracic gas volume and chestabdomen dimensions during anesthesia and muscle paralysis, Anesthesiology 55, 499–506, 1981. Jones, J.G., Faithfull, D., Jordan, C. and Minty, B., Rib cage movement during halothane anaesthesia in man, Br. J. Anaesth. 51, 399–407, 1979.
728
Warner
95.
Spens, H.J., Drummond, G.B. and Wraith, P.K., Changes in chest wall compartment volumes on induction of anaesthesia with eltanolone, propofol and thiopentone, Br. J. Anaesth. 76, 369–373, 1996. Morton, C.P. and Drummond, G.B., Change in chest wall dimensions on induction of anaesthesia: a reappraisal, Br. J. Anaesth. 73, 135–139, 1994. De Troyer, A. and Bastenier-Geens, J., Effects of neuromuscular blockade on respiratory mechanics in conscious man, J. Appl. Physiol. 47, 1162–1168, 1979. Kimball, W.R., Loring, S.H., Basta, S.J., De Troyer, A. and Mead, J., Effects of paralysis with pancuronium on chest wall statics in awake humans, J. Appl. Physiol. 58, 1638–1645, 1985. Saunders, N.A., Rigg, J.R., Pengelly, L.D. and Campbell, E.J., Effect of curare on maximum static PV relationships of the respiratory system, J. Appl. Physiol. 44, 589–595, 1978. Gal, T.J. and Arora, N.S., Respiratory mechanics in supine subjects during progressive partial curarization, J. Appl. Physiol. 52, 57–63, 1982. Smith, J.C. and Mead, J., Three degree of freedom description of movement of the human chest wall, J. Appl. Physiol. 60, 928–934, 1986. De Troyer, A., Estenne, M., Ninane, V., Van Gansbeke, D. and Gorini, M., Transversus abdominis muscle function in humans, J. Appl. Physiol. 68, 1010– 1016, 1990. Kaul, S.U., Heath, J.R. and Nunn, J.F., Factors influencing the development of expiratory muscle activity during anaesthesia, Br. J. Anaesth. 45, 1013–1018, 1973. Freund, F., Roos, A. and Dodd, R.B., Expiratory activity of the abdominal muscles in man during general anesthesia, J. Appl. Physiol. 19, 693–697, 1964. Taylor, A., The contribution of the intercostal muscles to the effort of respiration in man, J. Physiol. 151, 390–402, 1960. Warner, D.O. and Warner, M.A., Human chest wall function while awake and during halothane anesthesia, II. Carbon dioxide rebreathing, Anesthesiology 82, 20–31, 1995. Whitelaw, W.A. and Feroah, T., Patterns of intercostal muscle activity in humans, J. Appl. Physiol. 67, 2087–2094, 1989. De Troyer, A., Ninane, V., Gilmartin, J.J., Lemerre, C. and Estenne, M., Triangularis sterni muscle use in supine humans, J. Appl. Physiol. 62, 919–925, 1987. Drummond, G.B., Allan, P.L. and Logan, M.R., Changes in diaphragmatic position in association with the induction of anaesthesia, Br. J. Anaesth. 58, 1246–1251, 1986. Hedenstierna, G., Tokics, L., Lundquist, H., Andersson, T., Strandberg, A. and Brismar, B., Phrenic nerve stimulation during halothane anesthesia, Effects of atelectasis, Anesthesiology 80, 751–760, 1994. Drummond, G.B., Chest wall movements in anaesthesia, Eur. J. Anaesthesiol. 6, 161–196, 1989. Campbell, E.J.M., The role of the scalene and sternomastoid muscles in breathing in normal subjects, J. Anat. 89, 378–386, 1955.
96. 97.
98.
99.
100. 101. 102.
103.
104.
105. 106.
107. 108.
109.
110.
111. 112.
General Anesthesia and Respiratory Mechanics
729
113. Koepke, G.H., Smith, E.M., Murphy, A.J. and Dickinson, D.G., Sequence of action of the diaphragm and intercostal muscles during respiration, I. Inspiration, Arch. Phys. Med. Rehab. 39, 426–430, 1958. 114. Raper, A.J., Thompson, W.T., Jr., Shapiro, W. and Patterson, J.L., Jr., Scalene and sternomastoid muscle function, J. Appl. Physiol. 21, 497–502, 1966. 115. Tusiewicz, K., Bryan, A.C. and Froese, A.B., Contributions of changing rib cage-diaphragm interactions to the ventilatory depression of halothane anesthesia, Anesthesiology 47, 327–337, 1977. 116. Dempsey, J.A., Olson, E.B., Jr. and Skatrud, J.B., Hormones and neurochemicals in the regulation of breathing, in Handbook of Physiology, The Respiratory System. Mechanics of Breathing, Sec. 3, Vol. 3. Fishman, A.P., Macklem, P.T. and Mead, J., eds., Bethesda, MD, American Physiological Society, pp. 181–221, 1986. 117. Warner, D.O., Warner, M.A., Joyner, M.J. and Ritman, E.L., The effect of nitrous oxide on chest wall function in humans and dogs, Anesth. Analg. 86, 1058–1064, 1998. 118. Prabhakar, N.R., Mitra, J., Overholt, J.L. and Cherniack, N.S., Analysis of postinspiratory activity of phrenic motoneurons with chemical and vagal reflexes, J. Appl. Physiol. 61, 1499–1509, 1986. 119. Guedel, A.E., Inhalation anesthesia, A Fundamental Guide, New York, MacMillan, 1937. 120. Lumb, A.B., Petros, A.J. and Nunn, J.F., Rib cage contribution to resting and carbon dioxide stimulated ventilation during 1 MAC isoflurane anaesthesia, Br. J. Anaesth. 67, 712–721, 1991. 121. Estenne, M. and De Troyer, A., Relationship between respiratory muscle electromyogram and rib cage motion in tetraplegia, Am. Rev. Respir. Dis. 132, 53–59, 1985. 122. Warner, D.O., Warner, M.A. and Ritman, E.L., Human chest wall function during epidural anesthesia, Anesthesiology 85, 761–773, 1996. 123. Moote, C.A., Knill, R.L. and Clement, J., Ventilatory compensation for continuous inspiratory resistive and elastic loads during halothane anesthesia in humans, Anesthesiology 64, 582–589, 1986. 124. Iscoe, S.D., Central control of the upper airway, in Respiratory Function of the Upper Airway, Mathew, O.P. and Sant’Ambrogio, G., eds., New York, Marcel Dekker, Inc., pp. 125–192, 1988. 125. Rudomin, P., The electrical activity of the cricothyroid muscles of the cat, Arch. Int. Physiol. Biochem. 74, 135–153, 1966. 126. Sherrey, J.H. and Megirian, D., Spontaneous and reflexly evoked laryngeal abductor and adductor muscle activity of cat, Exp. Neurol. 43, 487–498, 1974. 127. Hershenson, M., Brouillette, R.T., Olsen, E. and Hunt, C.E., The effect of chloral hydrate on genioglossus and diaphragmatic activity, Pediatr. Res. 18, 516–519, 1984. 128. Hwang, J.C., St. John, W.M. and Bartlett, D., Jr., Respiratory-related hypoglossal nerve activity: influence of anesthetics, J. Appl. Physiol. 55, 785– 792, 1983.
730
Warner
129. Nishino, T., Kohchi, T., Yonezawa, T. and Honda, Y., Responses of recurrent laryngeal, hypoglossal, and phrenic nerves to increasing depths of anesthesia with halothane or enflurane in vagotomized cats, Anesthesiology 63, 404–409, 1985. 130. Nishino, T., Shirahata, M., Yonezawa, T. and Honda, Y., Comparison of changes in the hypoglossal and the phrenic nerve activity in response to increasing depth of anesthesia in cats, Anesthesiology 60, 19–24, 1984. 131. Ochiai, R., Guthrie, R.D. and Motoyama, E.K., Effects of varying concentrations of halothane on the activity of the genioglossus, intercostals, and diaphragm in cats: an electromyographic study, Anesthesiology 70, 812–816, 1989. 132. Rothstein, R.J., Narce, S.L., deBerry-Borowiecki, B. and Blanks, R.H., Respiratory-related activity of upper airway muscles in anesthetized rabbit, J. Appl. Physiol. 55, 1830–1836, 1983. 133. Drummond, G.B., Comparison of sedation with midazolam and ketamine: effects on airway muscle activity, Br. J. Anaesth. 76, 663–667, 1996. 134. Masuda, A., Ito, Y., Haji, A. and Takeda, R., The influence of halothane and thiopental on respiratory-related nerve activities in decerebrate cats, Acta. Anaesthesiol. Scand. 33, 660–665, 1989. 135. Horner, R.L., Innes, J.A., Holden, H.B. and Guz, A., Afferent pathway(s) for pharyngeal dilator reflex to negative pressure in man: a study using upper airway anaesthesia, J. Physiol. 436, 31–44, 1991. 136. Horner, R.L., Innes, J.A., Morrell, M.J., Shea, S.A. and Guz, A., The effect of sleep on reflex genioglossus muscle activation by stimuli of negative airway pressure in humans, J. Physiol. 476, 141–151, 1994. 137. Kuna, S.T., Inhibition of inspiratory upper airway motoneuron activity by phasic volume feedback, J. Appl. Physiol. 60, 1373–1379, 1986. 138. Kuna, S.T., Interaction of hypercapnia and phasic volume feedback on motor control of the upper airway, J. Appl. Physiol. 63, 1744–1749, 1987. 139. Onal, E., Lopata, M. and O’Connor, T.D., Diaphragmatic and genioglossal electromyogram responses to CO2 rebreathing in humans, J. Appl. Physiol. 50, 1052–1055, 1981. 140. Popovic, R.M. and White, D.P., Upper airway muscle activity in normal women: influence of hormonal status, J. Appl. Physiol. 84, 1055–1062, 1998. 141. Leiter, J.C., Knuth, S.L., Krol, R.C. and Bartlett, D., Jr., The effect of diazepam on genioglossal muscle activity in normal human subjects, Am. Rev. Respir. Dis. 132, 216–219, 1985. 142. Beydon, L., Goldenberg, F., Heyer, L., d’Ortho, M.P., Bonnet, F., Harf, A. and Lofaso, F., Sleep apnea-like syndrome induced by nitrous oxide inhalation in normal men, Respir. Physiol. 108, 215–224, 1997. 143. Drummond, G.B., Influence of thiopentone on upper airway muscles, Br. J. Anaesth. 63, 12–21, 1989. 144. Dhonneur, G., Combes, X., Leroux, B. and Duvaldestin, P., Postoperative obstructive apnea, Anesth. Analg. 89, 762–767, 1999. 145. Eastwood, P.R., Szollosi, I., Platt, P.R. and Hillman, D.R., Collapsibility of the upper airway during anesthesia with isoflurane, Anesthesiology 97, 786– 793, 2002.
General Anesthesia and Respiratory Mechanics
731
146. Horner, R.L., Innes, J.A., Murphy, K. and Guz, A., Evidence for reflex upper airway dilator muscle activation by sudden negative airway pressure in man, J. Physiol. 436, 15–29, 1991. 147. Wheatley, J.R. and Tangel, D.J., Mezzanotte, W.S., White, D.P., Influence of sleep on response to negative airway pressure of tensor palatini muscle and retropalatal airway, J. Appl. Physiol. 75, 2117–2124, 1993. 148. Mutoh, T., Kanamaru, A., Kojima, K., Nishimura, R., Sasaki, N. and Tsubone, H., Effects of volatile anesthetics on the activity of laryngeal ‘‘drive’’ receptors in anesthetized dogs, J. Vet. Med. Sci. 61, 1033–1038, 1999. 149. Nishino, T., Anderson, J.W. and Sant’Ambrogio, G., Effects of halothane, enflurane, and isoflurane on laryngeal receptors in dogs, Respir. Physiol. 91, 247–260, 1993. 150. Montravers, P., Dureuil, B. and Desmonts, J.M., Effects of i.v. midazolam on upper airway resistance, Br. J. Anaesth. 68, 27–31, 1992. 151. Leiter, J.C. and Daubenspeck, J.A., Selective reflex activation of the genioglossus in humans, J. Appl. Physiol. 68, 2581–2587, 1990. 152. van Lunteren, E. and Strohl, K.P., The muscles of the upper airways, Clin. Chest Med. 7, 171–188, 1986. 153. Hudgel, D.W., Martin, R.J., Johnson, B. and Hill, P., Mechanics of the respiratory system and breathing pattern during sleep in normal humans, J. Appl. Physiol. 56, 133–137, 1984. 154. Sauerland, E.K., Orr, W.C. and Hairston, L.E., DMG patterns of oropharyngeal muscles during respiration in wakefulness and sleep, Electromyogr. Clin. Neurophysiol. 21, 307–316, 1981. 155. Bartlett, D., Jr., Leiter, J.C. and Knuth, S.L., Control and actions of the genioglossus muscle, in Sleep and Respiration, Isa, F.G., Suratt, P.M. and Remmers, J.E., eds., New York, Wiley-Liss, Inc., pp. 99–108, 1990. 156. Hudgel, D.W. and Harasick, T., Fluctuation in timing of upper airway and chest wall inspiratory muscle activity in obstructive sleep apnea, J. Appl. Physiol. 69, 443–450, 1990. 157. Safar, P., Escarraga, L.A. and Chang, F., Upper airway obstruction in the unconscious patient, J. Appl. Physiol. 14, 760–764, 1959. 158. Nandi, P.R., Charlesworth, C.H., Taylor, S.J., Nunn, J.F. and Dore, C.J., Effect of general anaesthesia on the pharynx, Br. J. Anaesth. 66, 157–162, 1991. 159. Shorten, G.D., Opie, N.J., Graziotti, P., Morris, I. and Khangure, M., Assessment of upper airway anatomy in awake, sedated and anaesthetised patients using magnetic resonance imaging, Anaesth. Intensive Care. 22, 165–169, 1994. 160. Reber, A., Wetzel, S.G., Schnabel, K., Bongartz, G. and Frei, F.J., Effect of combined mouth closure and chin lift on upper airway dimensions during routine magnetic resonance imaging in pediatric patients sedated with propofol, Anesthesiology 90, 1617–1623, 1999. 161. Boidin, M.P., Airway patency in the unconscious patient, Br. J. Anaesth. 57, 306–310, 1985. 162. Mathru, M., Esch, O., Lang, J., Herbert, M.E., Chaljub, G., Goodacre, B. and vanSonnenberg, E., Magnetic resonance imaging of the upper airway, Effects
732
163.
164.
165.
166.
167.
168.
169.
170.
171.
172.
173.
174.
175.
Warner of propofol anesthesia and nasal continuous positive airway pressure in humans, Anesthesiology 84, 273–279, 1996. Abernethy, L.J., Allan, P.L. and Drummond, G.B., Ultrasound assessment of the position of the tongue during induction of anaesthesia, Br. J. Anaesth. 65, 744–748, 1990. Steinhart, H., Kuhn-Lohmann, J., Gewalt, K., Constantinidis, J., Mertzlufft, F. and Iro, H., Upper airway collapsibility in habitual snorers and sleep apneics: evaluation with drug-induced sleep endoscopy, Acta Otolaryngol. 120, 990–994, 2000. Litman, R.S., Hayes, J.L., Basco, M.G., Schwartz, A.R., Bailey, P.L. and Ward, D.S., Use of dynamic negative airway pressure (DNAP) to assess sedative-induced upper airway obstruction, Anesthesiology 96, 342–345, 2002. Pavlin, E.G., Holle, R.H. and Schoene, R.B., Recovery of airway protection compared with ventilation in humans after paralysis with curare, Anesthesiology 70, 381–385, 1989. Dhonneur, G., Lofaso, F., Drummond, G.B., Rimaniol, J.M., Aubineau, J.V., Harf, A. and Duvaldestin, P., Susceptibility to upper airway obstruction during partial neuromuscular block, Anesthesiology 88, 371–378, 1998. Hammer, J., Reber, A., Trachsel, D. and Frei, F.J., Effect of jaw-thrust and continuous positive airway pressure on tidal breathing in deeply sedated infants, J. Pediatr. 138, 826–830, 2001. Isono, S., Tanaka, A. and Nishino, T., Lateral position decreases collapsibility of the passive pharynx in patients with obstructive sleep apnea, Anesthesiology 97, 780–785, 2002. Meier, S., Geiduschek, J., Paganoni, R., Fuehrmeyer, F. and Reber, A., The effect of chin lift, jaw thrust, and continuous positive airway pressure on the size of the glottic opening and on stridor score in anesthetized, spontaneously breathing children, Anesth. Analg. 94, 494–499, 2002. Gunnarsson, L., Strandberg, A., Brismar, B., Tokics, L., Lundquist, H. and Hedenstierna, G., Atelectasis and gas exchange impairment during enflurane/nitrous oxide anaesthesia, Acta Anaesthesiol. Scand. 33, 629–637, 1989. Tokics, L., Hedenstierna, G., Strandberg, A., Brismar, B. and Lundquist, H., Lung collapse and gas exchange during general anesthesia: effects of spontaneous breathing, muscle paralysis, and positive end-expiratory pressure, Anesthesiology 66, 157–167, 1987. Strandberg, A., Tokics, L., Brismar, B., Lundquist, H. and Hedenstierna, G., Atelectasis during anaesthesia and in the postoperative period, Acta Anaesthesiol. Scand. 30, 154–158, 1986. Klingstedt, C., Hedenstierna, G., Lundquist, H., Strandberg, A., Tokics, L. and Brismar, B., The influence of body position and differential ventilation on lung dimensions and atelectasis formation in anaesthetized man, Acta Anaesthesiol. Scand. 34, 315–322, 1990. Hedenstierna, G., Atelectasis formation and gas exchange impairment during anaesthesia, Monaldi. Arch. Chest. Dis. 49, 315–322, 1994.
General Anesthesia and Respiratory Mechanics
733
176. Hachenberg, T., Lundquist, H., Tokics, L., Brismar, B. and Hedenstierna, G., Analysis of lung density by computed tomography before and during general anaesthesia, Acta Anaesthesiol. Scand. 37, 549–555, 1993. 177. Brismar, B., Hedenstierna, G., Lundquist, H., Strandberg, A., Svensson, L. and Tokics, L., Pulmonary densities during anesthesia with muscular relaxation—a proposal of atelectasis, Anesthesiology 62, 422–428, 1985. 178. Gunnarsson, L., Tokics, L., Gustavsson, H. and Hedenstierna, G., Influence of age on atelectasis formation and gas exchange impairment during general anaesthesia, Br. J. Anaesth. 66, 423–432, 1991. 179. Lindberg, P., Gunnarsson, L., Tokics, L., Secher, E., Lundquist, H., Brismar, B. and Hedenstierna, G., Atelectasis and lung function in the postoperative period, Acta Anaesthesiol. Scand. 36, 546–553, 1992. 180. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G. and Hedenstierna, G., Reexpansion of atelectasis during general anaesthesia may have a prolonged effect, Acta Anaesthesiol. Scand. 39, 118–125, 1995. 181. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G., Hogman, M. and Hedenstierna, G., Influence of gas composition on recurrence of atelectasis after a reexpansion maneuver during general anesthesia, Anesthesiology 82, 832–842, 1995. 182. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G., Reber, A. and Hedenstierna, G., Prevention of atelectasis during general anaesthesia, Lancet 345, 1387–1391, 1995. 183. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G., Reber, A. and Hedenstierna, G., Atelectasis and pulmonary shunting during induction of general anaesthesia—can they be avoided? Acta Anaesthesiol. Scand. 40, 524–529, 1996. 184. Reber, A., Engberg, G., Sporre, B., Kviele, L., Rothen, H.U., Wegenius, G., Nylund, U. and Hedenstierna, G., Volumetric analysis of aeration in the lungs during general anaesthesia, Br. J. Anaesth. 76, 760–766, 1996. 185. Neumann, P., Rothen, H.U., Berglund, J.E., Valtysson, J., Magnusson, A. and Hedenstierna, G., Positive end-expiratory pressure prevents atelectasis during general anaesthesia even in the presence of a high inspired oxygen concentration, Acta Anaesthesiol. Scand. 43, 295–301, 1999. 186. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G. and Hedenstierna, G., Airway closure, atelectasis and gas exchange during general anaesthesia, Br. J. Anaesth. 81, 681–686, 1998. 187. Hedenstierna, G., Tokics, L., Strandberg, A., Lundquist, H. and Brismar, B., Correlation of gas exchange impairment to development of atelectasis during anaesthesia and muscle paralysis, Acta Anaesthesiol. Scand. 30, 183–191, 1986. 188. Tokics, L., Strandberg, A., Brismar, B., Lundquist, H. and Hedenstierna, G., Computerized tomography of the chest and gas exchange measurements during ketamine anaesthesia, Acta Anaesthesiol. Scand. 31, 684–692, 1987. 189. Rothen, H.U., Sporre, B., Engberg, G., Wegenius, G. and Hedenstierna, G., Re-expansion of atelectasis during general anaesthesia: a computed tomography study, Br. J. Anaesth. 71, 788–795, 1993.
734
Warner
190. Rothen, H.U., Neumann, P., Berglund, J.E., Valtysson, J., Magnusson, A. and Hedenstierna, G., Dynamics of re-expansion of atelectasis during general anaesthesia, Br. J. Anaesth. 82, 551–556, 1999. 191. Benoit, Z., Wicky, S., Fischer, J.F., Frascarob, P., Chapuis, C., Spahn, D.R. and Magnusson, L., The effect of increased FIO(2) before tracheal extubation on postoperative atelectasis, Anesth. Analg. 95, 1777–1781, 2002. 192. Joyce, C.J. and Williams, A.B., Kinetics of absorption atelectasis during anesthesia: a mathematical model, J. Appl. Physiol. 86, 1116–1125, 1999. 193. Akca, O., Podolsky, A., Eisenhuber, E., Panzer, O., Hetz, H., Lampl, K., Lackner, F.X., Wittmann, K., Grabenwoeger, F., Kurz, A., Schultz, A.M., Negishi, C. and Sessler, D.I., Comparable postoperative pulmonary atelectasis in patients given 30% or 80% oxygen during and 2 hours after colon resection, Anesthesiology 91, 991–998, 1999. 194. Warner, D.O., Warner, M.A. and Ritman, E.L., Atelectasis and chest wall shape during halothane anesthesia, Anesthesiology 85, 49–59, 1996. 195. Rehder, K., Sessler, A.D. and Rodarte, J.R., Regional intrapulmonary gas distribution in awake and anesthetized-paralyzed man, J. Appl. Physiol. 42, 391–402, 1977. 196. Rehder, K., Knopp, T.J. and Sessler, A.D., Regional intrapulmonary gas distribution in awake and anesthetized-paralyzed prone man, J. Appl. Physiol. 45, 528–535, 1978. 197. Rodarte, J.R. and Fung, Y.C., Distribution of stresses within the lung, in Handbook of Physiology, The Respiratory System. Mechanics of Breathing, Sec. 3, Vol. 3. Fishman, A.P., Macklem, P.T., Mead, J. and Geiger, S., eds., Bethesda, MD, American Physiological Society, pp. 233–245, 1986. 198. Grassino, A.E. and Anthonisen, N.R., Chest wall distortion and regional lung volume distribution in erect humans, J. Appl. Physiol. 39, 1004–1007, 1975. 199. Roussos, C.S., Martin, R.R. and Engel, L.A., Diaphragmatic contraction and the gradient of alveolar expansion in the lateral posture, J. Appl. Physiol. 43, 32–38, 1977. 200. Grassino, A.E., Bake, B., Martin, R.R. and Anthonisen, R., Voluntary changes of thoracoabdominal shape and regional lung volumes in humans, J. Appl. Physiol. 39, 997–1003, 1975. 201. Hoppin, F.G., Jr., Green, I.D. and Mead, J., Distribution of pleural surface pressure in dogs, J. Appl. Physiol. 27, 863–873, 1969. 202. Rehder, K., Mallow, J.E., Fibuch, E.E., Krabill, D.R. and Sessler, A.D., Effects of isoflurane anesthesia and muscle paralysis on respiratory mechanics in normal man, Anesthesiology 41, 477–485, 1974. 203. Woo, S.W., Berlin, D. and Hedley-Whyte, J., Surfactant function and anesthetic agents, J. Appl. Physiol. 26, 571–577, 1969. 204. Enhorning, G., Pototschnik, R., Possmayer, F. and Burgoyne, R., Pulmonary surfactant films affected by solvent vapors, Anesth. Analg. 65, 1275–1280, 1986. 205. Paugam-Burtz, C., Molliex, S., Lardeux, B., Rolland, C., Aubier, M., Desmonts, J.M. and Crestani, B., Differential effects of halothane and thiopental on surfactant protein C messenger RNA in vivo and in vitro in rats, Anesthesiology 93, 805–810, 2000.
General Anesthesia and Respiratory Mechanics
735
206. Scheidt, M., Hyatt, R.E. and Rehder, K., Effects of rib cage or abdominal restriction on lung mechanics, J. Appl. Physiol. 51, 1115–1121, 1981. 207. Young, S.L, Tierney, D.F. and Clements, J.A., Mechanism of compliance change in excised rat lungs at low transpulmonary pressure, J. Appl. Physiol. 29, 780–785, 1970. 208. Warner, D.O., Vettermann, J., Brusasco, V. and Rehder, K., Pulmonary resistance during halothane anesthesia is not determined only by airway caliber, Anesthesiology 70, 453–460, 1989. 209. Joyner, M.J., Warner, D.O. and Rehder, K., Halothane changes the relationships between lung resistances and lung volume, Anesthesiology 76, 229–235, 1992. 210. Osmond, D.G., Functional anatomy of the chest wall, in The Thorax, Roussos, C. and Macklem, P.T., eds., New York, Marcel Dekker, Inc., pp. 199–233, 1985.
18 Recovery from Anesthesia
SHIROH ISONO
JACOB ROSENBERG
Chiba University Graduate School of Medicine Chiba, Japan
University of Copenhagen Hellerup, Denmark
I.
Introduction
The respiratory system responds to surgical stress and anesthetic interventions under influences of a variety of drugs used during the perioperative period in any surgical patient. A variety of respiratory complications may develop when the physiological responses fail to maintain respiratory homeostasis in patients at risk. Development of a respiratory complication is multi-factorial, and in turn, a risk factor can lead to a variety of respiratory complications. Some respiratory complications occur in patients without known risk factors, probably because of occasional failure to identify the risk factors. Depending upon the postoperative period, complication types vary as shown in Figure 18.1. In general, upper airway dysfunction and impairment of respiratory control occur in association with residual drug effects mostly on the day of the surgery. Constant hypoxemia due to impairment of lower airway and lung function usually peaks during days 1–3 after operation. Nocturnal oxygen desaturation, possibly related to alteration of postoperative sleep architecture, becomes evident during days 2–5 after operation. 737
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Figure 18.1 Schematic presentation of types of respiratory complications with associated pharmacological and physiological changes during the postoperative period. UA: upper airway; FRC: functional residual capacity; REM: rapid eye movement sleep.
In this chapter, we review current knowledge of physiological changes of the respiratory control systems and mechanisms of major respiratory complications occurring during the postoperative period. II.
Impairment and Recovery of Upper Airway Function after Anesthesia
A. Impaired Pharyngeal Function Clinical Significance of Pharyngeal Airway Obstruction During Recovery from Anesthesia
Pharyngeal obstruction is the most common respiratory complication during recovery from anesthesia [1,2]. Although changes of head or body positions or insertion of an artificial airway may resolve the problem in most cases, it is occasionally fatal because of development of severe hypoxemia causing dangerous cardiac arrhythmias and myocardial ischemia. Among those patients with risk factors for pharyngeal obstruction, patients with obstructive sleep apnea syndrome (OSAS) are to be managed very carefully because of their susceptibility to severe pharyngeal obstruction and cardiovascular morbidity [3]. This disorder has a high prevalence, affecting at least 4% of the adult male and 2% of the adult female population. Therefore, undiagnosed OSAS patients may frequently go to the operating room and postanesthesia care unit (PACU) [4]. Typical OSAS patients have
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all the risk factors for perioperative respiratory complications as identified by Asai et al. such as male sex, obesity and poor oropharyngeal view [2]. It is possible that most patients with respiratory complications during the early postoperative period have undiagnosed OSAS. Gupta et al. examined postoperative complications in OSAS patients undergoing hip or knee replacement and reported that the incidence of serious complications was significantly higher in untreated OSAS patients (21/68; 31%) compared with age-, BMI-matched non-OSAS subjects (9/101; 9%). Use of nasal CPAP prior to surgery, however, significantly reduced the complications in OSAS patients (3/33; 9%) [5]. Furthermore, Esclamado et al. reported severe perioperative complications including one death in OSAS patients undergoing surgical procedures on their upper airways (18/135; 13%) [6]. Seven of the eighteen cases had postoperative upper airway obstruction requiring emergent and appropriate interventions to the patients’ airways. These reports clearly indicate that pharyngeal obstruction occurring during recovery from anesthesia and surgery is more severe and more difficult to reverse by the patients themselves than preoperative pharyngeal obstruction occurring during sleep. Accordingly, anesthesia and surgery must have significant deleterious effects on airway maintenance mechanisms in patients already susceptible to pharyngeal obstruction. Mechanisms of Pharyngeal Obstruction
What are the mechanisms of deterioration of pharyngeal patency during recovery from anesthesia? While the mechanisms of upper airway obstruction are described in Chapter 10 in detail, Figure 18.2 summarizes the current understanding of how pharyngeal airway patency is maintained [7,8]. The pharyngeal airway size is determined by the balance between airway suction forces generated by contraction of the inspiratory pump muscles (i.e., the diaphragm, external intercostals) and the airway dilating forces generated by contraction of pharyngeal airway dilator muscles [9,10]. These two forces are precisely regulated by ‘neural mechanisms’ through the respiratory center of the medulla. Upper airway reflexes such as the negative pressure reflex [11–13], chemical stimuli such as hypoxemia [14] and hypercapnia [15], and the level of consciousness [16,17], all mutually interact and influence the respiratory center, thereby determining the balance. Another important factor in this balance model is the position of the fulcrum, i.e., the anatomy (structural mechanical properties) of the pharynx. Depending on the position, the pharyngeal airway size varies (‘anatomic mechanisms’). Isono et al. demonstrated that the position of the fulcrum (the anatomy) of OSAS patients is, graphically, to the right of normal subjects [18]. That is, OSAS patients have anatomical abnormalities of the pharynx.
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Figure 18.2 Balance of forces model for explanation of mechanisms of pharyngeal airway maintenance. Neural control mechanisms such as negative pressure reflex, chemical control, and consciousness level regulate the balance of the two antagonistic forces. Residual anesthetics, partial paralysis and upper airway receptor dysfunction could impair the neural mechanisms. Position of the fulcrum represents structural properties of the pharynx (anatomical mechanisms). Mechanical interventions shifting the fulcrum to the right (direction ‘a’) include mandibular advancement, neck extension, sniffing position, sitting position, and lateral position. Mechanical interventions shifting the fulcrum to the left (direction ‘b’) include neck flexion, mouth opening, and prone position.
A variety of mechanical interventions from outside the pharyngeal airway are known to alter the fulcrum position as shown in Figure 18.2 [19–24]. Accordingly, complicated but precise interaction between the neural and anatomical mechanisms maintains pharyngeal airway patency. Pharyngeal obstruction during recovery from anesthesia occurs when any part of the neural or anatomical mechanisms mentioned above is impaired. Pharyngeal obstruction usually occurs when patients lose consciousness or sleep under the influence of residual anesthetics, while severe pharyngeal obstruction, especially caused by marked pharyngeal swelling, is not always reversed even when patients are fully aroused. The sites of pharyngeal obstruction are not homogeneous among patients. In about one-half of the patients the retropalatal airway is exclusively obstructed, and in the remaining half both the retropalatal and retroglossal airways are obstructed [18].
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Contribution of Upper Airway Topical Anesthesia to Pharyngeal Obstruction
Superficially located upper airway receptors are shown to be responsible for the reflex augmentation of the upper airway muscle activity and for arousal response [11–13]. Upper airway topical anesthesia impairs the receptor function and therefore the neural mechanisms presented in Figure 18.2 occur. Increased frequency of apneas and increased inspiratory effort at termination of apnea during the night were reported in OSAS patients receiving oropharyngeal anesthesia [25]. Furthermore, Berry et al. demonstrated that upper airway topical anesthesia impaired the arousal response to airway occlusion [26]. Thus, upper airway anesthesia performed during the perioperative period possibly increases the risk of pharyngeal obstruction and prolongs the obstruction leading to severe hypoxemia, particularly in patients whose airway patency significantly depends upon these neural mechanisms. Contribution of Residual Anesthetics and Analgesia with Opioids to Pharyngeal Obstruction
Nishino et al. demonstrated that both inhalational and intravenous anesthetics decreased activity of the pharyngeal muscles more than of the inspiratory pump muscles [27], suggesting the development of pharyngeal narrowing during anesthesia in accordance with the balance of forces model (Figure 18.2). This differential effect persists during recovery from anesthesia, suggesting that postoperative recovery of breathing through an endotracheal tube does not assure unobstructed breathing after extubation. While the mechanisms of the selective depression of pharyngeal muscle activation are unclear, anesthetic drugs could directly depress the upper airway motor neurons, since the suppression was dose-dependent under constant chemical stimuli in their experiment. In addition, anesthetics could offset the threshold level of upper airway muscle activation in response to chemical stimuli like natural sleep, and the negative airway pressure reflex could be depressed by anesthetics, although these possibilities have not been tested. It should also be noted that the magnitude of differential suppression varied depending on the anesthetic drugs examined. Among these drugs, ketamine presented the least differential effects, indicating that ketamine has the advantage of maintaining a patent airway. Catley et al. described a high prevalence of pharyngeal obstruction events (obstructive apnea or paradoxic breathing in association with marked desaturation) within 16 h after surgery in patients receiving morphine for analgesia (456 episodes in 10 out of 16 patients). The use of regional analgesia resulted in less frequency of such events (0 episode in 16 patients) [28]. While they clearly mentioned that all the respiratory events occurred during sleep, it is unclear whether sleep architecture was similar between the
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groups. Accordingly, reduction of consciousness level due to opioid administration, or depressant effects of opioids on the respiratory centers, or interaction between these may be the key for understanding the mechanisms of this phenomenon. The level of consciousness is an important risk factor for the development of pharyngeal obstruction [29]. While general anesthesia evidently abolishes or suppresses the arousal response, the influences of low-dose or residual anesthetics on the arousal response to pharyngeal obstruction may have clinical significance, particularly during sleep shortly after recovery from anesthesia or under light sedation. Considering that the carotid body plays a significant role in the arousal response to hypoxia [30], and that a subanesthetic dose of halothane depresses the carotid body function [31], residual inhalational anesthetics may possibly inhibit the arousal response, and can lead to prolongation of apnea with development of severe hypoxemia and hypercapnia. In fact, inhalation of 0.2% of halothane decreased both the arousal threshold to eucapnic hypoxia and the hypoxic ventilatory response in sleeping dogs (Issa and Isono, unpublished observations). Anesthesiologists should recognize the clinical significance of the arousal response and the effects of anesthesia upon this response. Effects of Residual Muscle Paralysis on Pharyngeal Obstruction
Non-depolarizing muscle blockade has differential effects on various muscles [32,33]. As shown in Figure 18.3, the pharyngeal muscles are more sensitive to a small dose of vecuronium than is the diaphragm, and a low concentration of enflurane facilitates the differential effects of vecuronium [34]. Therefore, according to the balance of forces model, patients partially paralyzed with residual muscle blockade are at increased risk of pharyngeal airway obstruction, and the risk is further augmented under influences of both residual anesthetics and muscle blockade during recovery from anesthesia (Figure 18.3) [35]. As discussed above, the arousal response mediated through the carotid body plays a major role in termination of pharyngeal obstruction. Eriksson et al. found reduced hypoxic chemosensitivity in partially paralyzed humans [36]. Recently, the mechanisms of this observation were thoroughly explored by Igarashi et al., finding that vecuronium directly inhibits the carotid body neural response to hypoxia in a rat preparation [37]. Based on these findings, it would be reasonable to speculate that partial paralysis depresses arousal and may prolong the pharyngeal obstruction. Effects of Surgical and Anesthetic Interventions on Pharyngeal Obstruction
Extensive surgical procedures on the pharyngeal airway reportedly resulted in development of severe pharyngeal swelling followed by choking
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Figure 18.3 Changes in isometric contractility of the diaphragm (open circle) and geniohyoid muscles (open triangles) in response to intravenous injection of small doses of vecuronium in dogs anesthetized with pentobarbital alone (upper panel) and pentobarbital and 0.2 MAC of enflurane (lower panel). Differential effects between the muscles and augmentation of the differential effects by 0.2 MAC of enflurane are illustrated (Data from Ref. 35).
immediately after endotracheal extubation in OSAS patients [38,39]. Similarly, a high incidence of severe respiratory compromises (19/311, 6.1%), including one death and six re-intubations after anterior cervical spine surgery, was attributable to pharyngeal edema [40]. Even modest pharyngeal swelling caused by laryngoscopy and excessive fluid infusion during surgery may possibly have significant influences on pharyngeal airway maintenance immediately after surgery, although no study has examined this possibility. Damage to the mucosal receptors responsible for the pharyngeal negative airway pressure reflex may impair the neural mechanisms, and pharyngeal swelling itself may shift the fulcrum of the balance model to the right, increasing pharyngeal collapsibility. Furthermore, placement of a gastric tube through the nasal passage increases nasal resistance, possibly creating more negative suction forces at the collapsible pharynx. In addition, secretion retention due to inability to
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swallow during paralysis, and increase in secretions due to administration of neostigmine for reversal of muscle blockade, may increase surface adhesive force and thereby opening pressure of the pharyngeal airway, making the closed airway difficult to re-open [7,41,42]. All of these speculations need to be examined in future studies because of their potential clinical significance.
Effects of Body, Head and Neck Positions on Pharyngeal Obstruction
Body position is of great significance for pharyngeal airway patency in postoperative patients (Figure 18.2). Due to the presence of a variety of drainage tubes or protection of the operative wound, postoperative patients are often lying on their backs. If possible, the lateral and sitting positions are advantageous for upper airway maintenance because of less gravitational effects on the upper airway in these body positions [21,43]. In this context, the prone position is expected to have the smallest gravitational impact on the airway, leading to improvement of pharyngeal airway patency. However, Safar et al. demonstrated that there was no improvement of tidal volume in the prone position when compared with the supine [44]. In agreement with this, Ishikawa et al. recently reported that prone positioning did not improve pharyngeal collapsibility, but rather increased the closing pressure of the passive pharynx in normal infants under general anesthesia with muscle paralysis to the equivalent level of patients with OSAS [22]. While these observations are interesting and the underlying mechanisms remain unclear, the results strongly suggest a significant role for neural control mechanisms in compensation for structural pharyngeal narrowing, particularly in infants. Since residual anesthetics possibly impair many neural reflexes including the arousal response, which is crucial for recovery from pharyngeal obstruction and from any life-threatening choking event, prone positioning should be avoided, particularly in small children after surgery. Use of a pillow during recovery from anesthesia is controversial. Head elevation with a pillow produces neck flexion at the lower cervical spine and extension at the upper cervical spine as long as the face is maintained straight up. This so-called ‘‘sniffing position,’’ which is a standard head position during induction of anesthesia, improves airway patency [45]. However, it is difficult to maintain the face straight up with head elevation, and the neck is often flexed at the higher cervical spine, resulting in narrowing of the pharyngeal airway [46]. Therefore, it is recommended that a pillow be used during emergence from anesthesia and at extubation of an endotracheal tube, but a high pillow should not be used in the PACU when
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there is less observation and residual anesthetics may still impair the neural compensatory mechanisms. B. Impaired Laryngeal Airway Maintenance Function Laryngospasm During Emergence from Anesthesia
The physiological basis of laryngospasm is reviewed in detail in Chapter 7. In short, laryngospasm is defined as a glottic closure induced by afferent inputs from airway receptors. Physiologically, this reflex functions to prevent foreign material entering the lower airways. It is, however, a clinically detrimental reflex leading to severe hypoxemia, and is recognized as a respiratory complication commonly observed immediately after tracheal extubation, particularly in children. Prevalence of laryngospasm during anesthesia is reported to be about 1% [47]. Half of the anesthesiarelated laryngospasm cases requiring treatment occurred after tracheal extubation in children [48]. Documented risk factors for laryngospasm include light anesthesia level [49,50], younger children [50], use of isoflurane [49], airway surgery [48,50], and upper respiratory tract infection [49,51]. It should be noted that the latter three risk factors are related to presence of irritants or irritating drug on the airway mucosa. Pounder et al. reported that tracheal extubation immediately after emergence from halothane or isoflurane anesthesia was associated with significantly more episodes of oxygen desaturation than tracheal extubation under deep anesthesia in 1- to 4-year-old children, while the difference was more apparent in children anesthetized with isoflurane than those anesthetized with halothane [49]. In another study, however, the frequency of laryngospasm did not differ between tracheal extubation during deep anesthesia and that after emergence in children anesthetized with halothane [51]. Complete laryngeal closure can last for several minutes, and many case reports have documented development of severe hypoxemia and ‘negative pressure pulmonary edema’ after post-extubation laryngospasm [52]. Laryngospasm may be effectively prevented by intravenous administration of lidocaine, although this is still controversial [53,54]. Although application of positive pressure is believed to be effective to resolve the laryngospasm, there is no systematic study evaluating its effectiveness. Administration of muscle blockers such as succinylcholine is the most effective treatment of laryngospasm, while intravenous administration of anesthetics such as propofol is effective in some cases [55]. Laryngeal Edema and Stridor Immediately after Extubation
Interaction between an endotracheal tube and the larynx during anesthesia and surgery significantly influences breathing after tracheal
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Flow (liters · s−1) Figure 18.4 Curves show translaryngeal pressure–flow relationships obtained before and after minor surgery in a patient whose trachea was intubated with an endotracheal tube for approximately 3 h. Note significant increase in the laryngeal resistance after endotracheal intubation (Data from Ref. 58).
extubation [56,57]. As illustrated in Figure 18.4, Tanaka et al. demonstrated that placement of an endotracheal tube even for 3-4 h significantly increased laryngeal resistance in association with development of asymptomatic but substantial laryngeal edema [58,59]. The increase in laryngeal resistance and edema formation was less severe in patients with laryngeal mask airway placement than those with endotracheal tube placement [59]. The standard endotracheal tube size used during anesthesia is larger than the normal glottic width of 6–8 mm in adults, suggesting that direct contact forces by the endotracheal tube could cause trauma and edema formation at the vocal cords [57]. Weymuller et al. found that the contact pressure could exceed 200 mmHg, far greater than the mucosal capillary perfusion pressure of 30 mmHg [60]. In a dog preparation, tracheal intubation with an oversized tube for 8 h resulted in mucosal ischemia and necrosis with tissue edema, particularly around the arytenoids [57]. Laryngeal edema due to placement of an endotracheal tube develops in the supra- and subglottic regions in addition to the vocal cords. The subglottic region is considered to be narrowest in neonates and small children, while recent MRI analysis indicates the narrowest region to be at the vocal cords, even in small children during wakefulness [61]. Even a mild level of edema formation at the subglottic region in small children results in a marked reduction of cross-sectional area and therefore increases laryngeal resistance.
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In fact, the risk factors for development of laryngeal edema after tracheal extubation include use of a tight-fitting tracheal tube, younger children, trauma at tracheal intubation, duration of intubation, and coughing during intubation [62]. The cuff-leak test, in which air leakage is detected around the tube with the cuff deflated and with the tube occluded, is recommended as a simple and useful method for diagnosis of severe laryngeal edema and for prediction of successful extubation [63,64]. Intravenous administration of dexamethasone decreases the need for reintubation in neonates after mechanical ventilation, while the effectiveness of nebulized adrenaline on the laryngeal edema is controversial [65,66]. Regardless of the frequency of laryngeal edema after surgery, postoperative laryngeal stridor infrequently occurs immediately after emergence from anesthesia whereas postextubation stridor occurs in 4–18% of patients after prolonged intubation in the intensive care unit [67,68]. Stridor develops due to severe but incomplete laryngeal narrowing during spontaneous breathing, even in awake subjects, within 6 h after tracheal extubation [69]. Stridor should be treated, although development of hypoxemia is easily prevented by supplementary oxygen administration. Increased inspiratory drive due to hypercapnia increases the work of breathing and some cases require reintubation, tracheostomy and prolonged intensive care [70]. Stridor results from interaction between the gas traveling through the airway and narrowed laryngeal airway wall during inspiration, producing vibration of the vocal cords with a high-pitched characteristic sound different from the snoring sound [71]. The velocity of the air at the passively narrowed vocal cords increases during inspiration, converting potential energy to kinetic energy. Due to the increased kinetic energy, lateral wall pressure at the vocal cords decreases and, therefore, narrows the vocal cords further during inspiration. Fiberoptic examination reveals paradoxical vocal cord movement [69]. While this passive mechanism is generally believed to be responsible for the postoperative stridor, no physiologic measurement has been systematically performed because of difficulty in predicting the occurrence of postoperative stridor in a particular patient. Recently, Isono et al. found inspiratory activation of laryngeal adductor muscle in association with the development of stridor in patients with multiple system atrophy [72]. They suggested imbalance between forces generated by vocal cord adductor muscle contraction and those generated by abductor muscle contraction (active mechanism) as a possible mechanism for the generation of stridor. Application of CPAP eliminated the stridor and inspiratory adductor muscle activation, strongly suggesting that the inspiratory adductor muscle activity is induced by a negative pressure reflex; CPAP functions to depress the reflex in addition to an airway splint effect. In this context, effectiveness of benzodiazepines for elimination of postoperative laryngeal stridor may be explained by reduction of negative
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inspiratory pressure [69]. Future studies should examine whether the active mechanism plays a role in the development of postoperative and/or postextubation stridor and whether CPAP application is effective for treatment of the stridor.
C. Impaired Airway Protective Functions
Aspiration pneumonia is a life-threatening major complication after surgery, and impaired airway protective mechanisms can be responsible (see Nishino, Chapter 7, this volume). Postoperative laryngeal incompetence is common, particularly in patients managed with endotracheal tubes during anesthesia, although it does not result in clinical problems such as aspiration pneumonia in most cases. In earlier investigations, 11–33% of postoperative patients aspirated radiopaque dye into the lower airways upon swallowing the dye immediately after extubation [73,74]. Furthermore, Burgess et al. noted a time-dependence in the incidence of radiopaque aspiration in patients undergoing cardiac surgery (33% immediately after extubation, 20% 4 h after extubation, and 5% 8 h after extubation) [74]. In contrast to previous studies, a recent investigation reported the incidence to be around 5% [75]. Although reduction of the incidence can be explained by improvement in patient management, such as the use of less irritating material for tracheal tubes with high volume and low pressure cuffs, and short-acting anesthetics and muscle blockers, this does not indicate complete recovery of airway defensive mechanisms immediately after surgery. Interestingly, aspiration occurred during wakefulness without signs of laryngeal irritations (e.g., cough). While reduction of consciousness level is reported to depress the swallowing reflex [76] and alter the pattern of laryngeal airway reflexes [77], the consciousness level is unlikely to play a major role in the mechanisms of postextubation aspiration. Swallowing dysfunction possibly contributes to aspiration. Although no study has systematically examined changes in swallowing function after surgery, one possible mechanism is dysfunction of the pharyngeal receptor eliciting the swallowing reflex, due to development of pharyngeal edema. Alternatively, partial paralysis due to residual muscle blockade may impair swallowing function as demonstrated experimentally [78–80]. Lack of laryngeal irritation manifested by cough reflex further suggests depression of laryngeal airway defensive reflexes after extubation. Postoperative laryngeal edema possibly damages mucosal receptors for the laryngeal reflexes, depressing the reflexes. Hasegawa and Nishino recently suggested the contribution of a central adaptation mechanism to alteration of upper airway reflex patterns during emergence from anesthesia (Figure 18.5) [81]. Future studies need to evaluate upper airway receptor function for induction of airway defensive reflexes after surgery.
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Figure 18.5 Difference in airway reflexes elicited while decreasing sevoflurane concentration (ETsevo) before and after surgery is shown. Increase in respiratory frequency resulted in increase in minute ventilatory volume and decrease in end-tidal CO2 concentration (ETCO2 ) after surgery. Apnea with swallowing reflexes indicated by arrows was observed during the post-surgical period whereas apnea and a series of expiration reflexes were evident during the pre-surgical period in this patient. ETsevo at the occurrence of airway reflexes did not differ from that before or after surgery. EMG: electromyogram; airflow: airflow measured by a pneumotachograph; in: inspiration; ex: expiration; VT: tidal volume; PAW: airway pressure (Data from Ref. 81).
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Impairment and Recovery of Chemical Control of Breathing after Anesthesia
A. Postanesthetic Residual Drug Effects on Control of Breathing
Residual inhalation anesthetics and muscle blockades after surgery influence chemical control of breathing (Figure 18.6). Knill and other researchers (see Dahan et al., Chapter 16, this volume) consistently found that low concentrations of inhalational anesthetics, less than MAC-awake, significantly depress hypoxic ventilatory response in humans whereas hypercapnic ventilatory response is preserved in such conditions [31]. Depression of carotid chemosensitivity with small doses of halothane was confirmed as the mechanism for the reduced hypoxic ventilatory response in animal experiments [82,83]. Partial paralysis with vecuronium is also reported to depress hypoxic ventilatory response in human volunteers [36] and to inhibit neurotransmission of the rat carotid body [37]. Accordingly, carotid body function may be significantly impaired in patients during recovery from inhalational anesthesia combined with muscle blockade. No doubt, absence of increased ventilation during hypoxemia is of clinical significance. Hypoxemia due to central hypoventilation primarily caused by
in ase e Incre tory driv a r i p s re R HCV R HV
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Figure 18.6 Possible impairment of chemical control of breathing during the immediate postoperative period is illustrated. Residual inhalation anesthetics significantly impair the regulatory effects indicated by the open arrows. Use of opioids predominantly depresses the hypercapnic ventilatory response indicated by the closed arrows. Note that use of opioids immediately after anesthesia with inhalation anesthetics possibly results in loss of all respiratory control mechanisms including arousal response and dypneic sensation even when severe hypoxemia develops. HVR: hypoxic ventilatory response; HCVR: hypercapnic ventilatory response; CNS: central nervous system.
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depression of hypercapnic ventilatory response may be reversed by preservation of the ‘back-up’ hypoxic ventilatory response. However, it should be noted that increased ventilatory drive itself does not usually reverse the hypoxemia resulting from upper airway obstruction and impaired lung mechanics. Knill and Gelb compared the carotid body to a watchdog, sensing hypoxemia and activating the alarm [84]. Increased ventilation, dyspneic sensation and arousal are considered to be significant clinical signs and survival responses produced by carotid body activation. Clinicians should recognize the possible depression of these alarm functions of the carotid body in patients during recovery from anesthesia. B. Opioids and Control of Breathing
Opioids are known to decrease the slope of hypercapnic response while increasing the apneic threshold. Absence or depression of both hypoxic and hypercapnic responses is a life-threatening condition. Baraka reported two illustrative cases following excision of carotid body tumors [85]. The patients developed severe hypoxemia with marked respiratory depression in association with opioid administration after surgery. Opioid-induced respiratory depression in the absence of peripheral chemoreceptor drive contributed to the respiratory complication. It should be noted that a similar condition is possibly produced in patients receiving opioids immediately after inhalational anesthesia. Furthermore, recent investigations clearly demonstrated that opioids significantly depress hypoxic ventilatory response as well as hypercapnic response in humans [86,87]. Both animal [88] and human [87] studies indicate that the effect of morphine on hypoxic ventilatory response is likely to be mediated centrally rather than peripherally. Delayed respiratory depression is reported to occur in the ward even after confirming stable respiration in patients anesthetized with opioids [89,90]. Decrease in environmental stimulation after postoperative intensive nursing care could unmask an opioid-induced, long-lasting depressant effect on ventilation. Alternatively, biphasic pharmacokinetic characteristics of fentanyl could explain the phenomenon [91]. Becker et al. found recurrence of depression in the hypercapnic response in patients anesthetized with either fentanyl or fentanyl–droperidol after initial complete recovery from the hypercapnic response [89]. Interestingly, combined use of droperidol with fentanyl appeared to decrease the respiratory depressant effects of fentanyl. However, these observations were not confirmed by a later study presenting progressive recovery of the hypercapnic response [92]. C. Effects of Postoperative Pain on Control of Breathing
Postoperative residual drug effects on control of breathing interact with surgical pain. Lam et al. examined effects of surgical stimulation on
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breathing in enflurane-anesthetized humans [93]. They found a significant increase in ventilation with reduction of PaCO2, without changing both hypercapnic and hypoxic ventilatory responses. Nishino and Kochi [94] found that, in sevoflurane-anesthetized human, the apneic threshold for carbon dioxide with surgical stimulation was reduced. Sarton et al. further demonstrated that experimentally induced acute pain increased resting ventilation without affecting the hypoxic ventilatory response significantly depressed by low-dose sevoflurane [95]. Similarly, Borgbjerg et al. found that experimental pain attenuated morphine-induced respiratory depression while increasing the ventilation intercept (at 7.2% inspired CO2) of the hypercapnic response curve without changing the slope [96]. All these studies strongly suggest that acute surgical pain does not normalize the hypoxic and hypercapnic responses depressed by postoperative residual drug effects, although it increases resting ventilation. IV.
Impairment and Recovery of Lung Function after Surgery
A. Lung Volume Changes and Constant Hypoxemia after Surgery
As extensively reviewed by Warner in Chapter 17 of this volume, pulmonary atelectasis identified by CT-scan develops in dependent lung regions shortly after induction of general anesthesia, resulting in reduction of functional residual capacity (FRC) by approximately 500 ml and impairment of gas exchange [97]. Cephalad shift of diaphragm position at FRC [98], possibly resulting from reduction of tonic activity of the diaphragm [99], may produce atelectasis during anesthesia. After upper abdominal surgery, FRC further decreases and nadirs on the first or second postoperative day (70% of preoperative FRC) in accordance with maximum reduction of PaO2 [100–102], whereas postoperative FRC reduction is small and not consistently observed in patients after superficial or extremity surgery [100,103]. Compared with open laparotomy for cholecystectomy, laparoscopic surgery results in less reduction of FRC and PaO2, which is furthermore reversed within 24 h [104]. In obese patients, however, this is not the case (Figure 18.7). Percentage of pulmonary atelectasis at dependent lung regions was shown to progressively increase even after laparoscopic surgery in morbidly obese patients [105]. Mechanisms of the FRC reduction after surgery appear to differ from those during general anesthesia. Recent evidence strongly suggests that inhibitory reflexes of phrenic nerve activity arising from abdominal wall and/or viscera contribute to the FRC reduction [106,107]. Ford et al. assessed breathing and diaphragm function in patients undergoing cholecystectomy [106]. Breathing pattern was characterized by
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Figure 18.7 Images of computed tomography taken before induction of anesthesia, immediately after extubation and 24 h later after laparoscopic cholecystectomy in a morbidly obese patient (upper three images) and a non-obese patient (lower three images). The slices were realized at the level of the interventricular septum. Note that significant atelectasis at the dorsal region is evident even 24 h after surgery in the obese patient (Data from Ref. 105).
rapid shallow breaths, predominantly from rib cage movement. Significant decrease in transdiaphragmatic pressure indicating inhibition of diaphragm contraction was suggested to be responsible for the observed respiratory pattern and reduction of FRC. Dureuil et al. further suggested that inhibitory reflexes arising from peripheral stimuli possibly cause the postoperative dysfunction because the diaphragm contractility did not change during bilateral phrenic nerve electrical stimulation [107]. While the afferent pathway of the reflex arc has not been determined yet, splanchnic and peritoneal irritation produced by the surgery is speculated to induce the inhibitory reflexes based on the findings of inhibition of diaphragm contraction by esophageal and bowel distension [108,109], as well as by gallbladder stimulation [110]. In an early study, Spence and Smith demonstrated in patients after upper abdominal surgery that thoracic epidural blockade with bupivacaine resulted in better oxygenation and less pulmonary complications than intramuscular administration of morphine [111]. Results of this pioneering study suggest a role for pain in the postoperative diaphragm dysfunction and possibly the spinal reflex arc of the inhibitory phrenic activity.
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However, recent studies found that relief of pain by thoracic epidural blockade only minimally reverses the diaphragm dysfunction and FRC reduction after abdominal surgery [112,113]. A more recent study failed to find superiority of epidural analgesia using bupivacaine and opioids over parenteral opioids in terms of postoperative pulmonary complications, despite the significantly better pain relief in patients receiving epidural analgesia [114]. B. Respiratory Ciliary Dysfunction after Surgery
Mucous transport is an important defense mechanism against respiratory tract infections. Impairment of mucous transport may contribute to postoperative pulmonary complications. In fact, Gamsu et al. found significant association between prolonged impairment of ciliary function and development of radiographically visible atelectasis in patients undergoing upper abdominal surgery [115]. Patients undergoing lower extremity surgery whose ciliary function progressively recovered within 48 h after surgery did not experience any pulmonary complications. Mechanisms of the prolonged ciliary dysfunction after abdominal surgery are unclear at present. Inhalational anesthetics such as halothane, enflurane and isoflurane depressed respiratory ciliary function by 20–25% in in vitro preparations [116,117]. Raphael et al. further observed recovery of ciliary beat frequency within 3 h after exposure to inhalational anesthetics [118]. In contrast, intravenous anesthetics such as midazolam, fentanyl, and propofol (but not thiopental) did not alter respiratory ciliary beat frequency [119–121]. Accordingly, anesthesia per se may not be a primary cause of postoperative ciliary dysfunction. Insufficient humidity of inspired gases [122], mechanical ventilation [123], placement of an endotracheal tube [124,125] and aging [126] have been reported to cause ciliary dysfunction. More importantly, smokers had a slower bronchial mucus transport than non-smokers during anesthesia and experienced a higher incidence of pulmonary complications after abdominal surgery [127]. However, none of the possibilities mentioned above explains preferential occurrence of prolonged ciliary dysfunction in patients undergoing abdominal surgery. C. Postoperative Atelectasis and Pneumonia
In clinical practice, the constant hypoxemia associated with FRC reduction after surgery is well managed in most cases by administration of oxygen. There is no doubt that FRC reduction contributes to postoperative pulmonary complications such as radiographically visible atelectasis and pneumonia. However, additional risk factors may be necessary for the complications to develop. A recent prospective study conducted by BrooksBrunn identified age 460 years, BMI 427 kg/m2, impaired cognitive
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function, history of cancer, smoking history in past 8 weeks, and upper abdominal incision as risk factors for development of postoperative pulmonary complications, which occurred in 22.5% (126/561) of patients after abdominal surgery [128]. Although pulmonary function tests can be useful for optimizing preoperative management of preexisting lung disease, no variables obtained from the pulmonary function tests have succeeded in predicting postoperative pulmonary complications [129]. Prophylactic mechanical ventilation in high-risk patients did not result in improvement of oxygenation and reduction of pulmonary complications [130]. Because reduction of FRC is a fundamental mechanism of pulmonary atelectasis and pneumonia after upper abdominal surgery, prevention and/or treatment of the pulmonary complications should be directed to increase FRC. Intermittent voluntary deep inspiration [131] and sitting position improve oxygenation through an increase in FRC. Currently available strategies for reduction of postoperative pulmonary complications include respiratory physiotherapy [132], incentive spirometry [133], intermittent positive-pressure breathing (IPPB) [134] and periodic application of CPAP [135]. While each of these was equally effective, studies failed to show an additive effect for any of these techniques [136]. While promising results have been reported in pharmacological therapies with doxapram [137], aminophylline [138], amrinone [139], and olprinone [140], future studies need to re-evaluate the efficacy of the treatments and clinical outcomes. V.
Late Postoperative Nocturnal Hypoxemia
A. Clinical Significance of Late Postoperative Hypoxemia Nature of Late Postoperative Nocturnal Hypoxemia
Until recently, the importance of the late postoperative period in the development of disordered breathing during sleep, resulting in episodic hypoxemia, was unclear. Adequate oxygenation, diagnosed by blood-gas analysis during wakefulness, does not assure stable oxygenation during sleep in this period. Continuous measurement of arterial oxygen saturation with a pulse oximeter after surgery revealed development of nocturnal episodic hypoxemia [141,142]. This late postoperative hypoxemia is characterized by episodic and periodic occurrences of hypoxemia during sleep in postoperative patients breathing room air. The periodicity and the severity of the episodic desaturation have been characterized using oximetric variables such as oxygen desaturation index (ODI, h 1) which is usually defined as the number of oxygen desaturation episodes exceeding 4% or more below the baseline per hour, the percent of time spent at SpO2 5 90% (CT90, %), the lowest SpO2 value, and mean of the SpO2 values [143,144]. A 2-min window is usually set for the determination of the desaturation episode.
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However, we lack full knowledge of disordered breathing patterns during episodic hypoxemia due to lack of polysomnographic evaluation during late postoperative nights. Using a portable monitoring system, Rosenberg et al. reported that apneas associated with oxygen desaturation were not central and were either obstructive or mixed, while 23% of hypopneas also led to desaturation [145]. This observation strongly suggests that the pattern of disordered breathing during episodic hypoxemia is obstructive in nature. However, the nature of the hypopneic events was not determined because the portable monitoring system did not allow further characterization of the hypopneas. Full polysomnographic assessment should be performed in future studies to clarify the pattern of disordered breathing during late postoperative nights. While no large-scale survey of late postoperative hypoxemia has been conducted, Rosenberg et al. demonstrated that 23% of patients (18/78) had more than 30 episodes of desaturation during the second postoperative night after major abdominal surgery without oxygen therapy, and 38% of the patients (30/78) had more than one episode of sudden desaturation to below 80% SpO2 [146]. Using generally accepted criteria for abnormal oxygenation during sleep in adults (ODI greater than 5 h 1 and CT90 less than 1%), the incidence of late postoperative desaturation was 50% in the study by Reeder et al. [147]. The prevalence of late postoperative desaturation appears to be greater than that of sleep disordered breathing (SDB) in the general adult population (AHI greater than 5 h 1 was found in 23% of adult males and 9% of adult females) [4]. Most studies examining nocturnal oxygenation after surgery without oxygen therapy indicate the presence of deterioration of nocturnal oxygenation on the second or third postoperative night when compared with preoperative oxygenation [141,148,149]. Furthermore, a study examining the time course of nocturnal oxygenation after major surgery for up to five nights clearly demonstrated that the episodic hypoxemia during sleep was most frequent on the third postoperative night and oxygenation on the fifth night did not differ from preoperative oxygenation [150]. While there is no doubt that the presence of preoperative SDB is a major and significant risk factor for late postoperative desaturation [151,152], the higher prevalence of this condition, deterioration of nocturnal oxygenation and recovering from the hypoxemia after surgery, all strongly suggest that late postoperative nocturnal desaturation can develop even in subjects without preoperative SDB. Clinical Significance of Late Postoperative Nocturnal Hypoxemia
Episodic hypoxemia during sleep is very common, and occurs every night in patients with OSA without apparent acute detrimental effects in most of these patients. While this may indicate clinical unimportance for episodic hypoxemia after surgery, one should recognize that, in addition to increases
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in oxygen demand under surgical stress, oxygen supply decreases due to the deterioration of nocturnal oxygenation after surgery, resulting in tissue hypoxemia, and additionally, in impaired function of important organs, particularly in patients with risk factors for this dysfunction. While no study has clearly demonstrated direct association between late postoperative nocturnal hypoxemia and morbidity and mortality in postoperative patients, episodic hypoxemia could be responsible for negative clinical outcomes in any vital organ. Perioperative myocardial infarction is a major cause of death associated with anesthesia and surgery. Myocardial ischemia and infarction occur in patients with ischemic heart disease, with a peak incidence intraoperatively and on the third postoperative day when episodic hypoxemia most commonly develops [153,154]. Rosenberg et al. demonstrated a temporal relationship between episodic hypoxemia and myocardial infarction and arrhythmias in the late postoperative period [148], confirmed by other researchers [155,156], and also in non-surgical patients with acute myocardial infarction [157]. Periodic swings of heart rate and blood pressure typically seen in OSA episodes also increase oxygen demand at a time of decreased oxygen supply, shifting the oxygen balance to tissue hypoxia. The tissue hypoxia may be critical, particularly when the myocardium is already ischemic due to coronary arterial disease in postoperative patients with tachycardia, developing severe myocardial ischemia. Oxygen therapy appears to be beneficial, increasing oxygen saturation and decreasing heart rate [158]. Because severe hypoxemia can cause dangerous arrhythmias such as ventricular tachycardia and atrio-ventricular block [148,159], one could hypothesize that episodic hypoxemia might be a pathogenic factor for unexpected postoperative death. In support of this hypothesis, Rosenberg et al. found that approximately two-thirds of totally unexpected postoperative deaths occurred at night [160]. Thus, a single episode of severe hypoxemia could lead to a fatal outcome. In addition, after major abdominal surgery, patients were proposed to have increased sympathetic tone with lack of circadian variation [161]. This increased sympathetic tone could peak when severe episodic hypoxemia occurs during REM sleep [142,149]. This is an interesting finding because SIDS victims had a high peak of sympathetic tonus in the late hours of night when most SIDS deaths occur [162]. Postoperative delirium is more common in the elderly, with an incidence of 5–10% after major surgery and peaking in the middle of the first week after surgery [163,164]. The risk factors generally identified include age, preoperative dementia, severe illness, metabolic and electrolyte imbalance, use of alcohol and psychoactive drugs, and infections [165,166], whereas type of anesthesia [167] and postoperative analgesia [168] appear to be of less importance. Hypoxemia was reported to be a significant risk factor for mental confusion immediately after surgery [169], and delirium occurring on the second postoperative day [164] as also reported in
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non-surgical patients [170]. Rosenberg and Kehlet further found that mental function on the third day after laparotomy correlated with mean oxygen saturation on the preceding night, and there was no relationship between mental function and other perioperative variables such as age, duration of surgery and postoperative doses of opioids [171]. Furthermore, significance of nocturnal hypoxemia in postoperative delirium was supported by the successful treatment of postoperative delirium by simple oxygen administration [167,172]. Despite antibiotic prophylaxis, wound infection rates are still about 2–10% in major surgeries [173]. Reduced oxygen supply to the surgical wound impairs healing and lowers resistance against bacterial wound infection in experimental and clinical studies [174–176]. Postoperative treatment with oxygen usually produces a significant increase in arterial as well as in subcutaneous oxygen tension [174,177–179]. Accordingly, observations made during the second and third postoperative nights strongly suggest detrimental effects of postoperative nocturnal hypoxemia on wound healing processes and on resistance against bacterial wound infections. Unfortunately, no study has critically tested this possibility. B. Postoperative Changes in Sleep Architecture Sleep Fragmentation and REM Rebound after Surgery
All studies examining changes in postoperative sleep architecture have shown suppression of REM sleep during the immediate postoperative night and a rebound increase in REM sleep during the third and forth postoperative nights [142,149,180]. Knill et al. examined sleep architecture before and after surgery in detail [142] (Figure 18.8). Sleep during the first and second postoperative nights was characterized by sleep fragmentation with reduction of both slow-wave sleep and REM sleep in addition to reduction of total sleep time. By contrast, they observed ‘rebound’ increases in slow-wave sleep and REM sleep during the third and fourth postoperative nights. Interestingly, the REM rebound was mainly due to an increase in ‘phasic REM’, which was confirmed by frequent eye movements and was accompanied by frequent nightmare reports. Surges of heart rate with increased cardiac sympathetic activity were reported during the ‘phasic REM’ period in dogs [181]. Respiration is highly irregular during REM sleep, and obstructive as well as central apnea/hypopnea with mild oxygen desaturation can develop due to loss of upper airway muscle activity and inhibition of primary respiratory muscle activity even in normal subjects. REM sleep is often the time when a patient with OSAS will have the most significant apneas and desaturations. Rosenberg et al. reported the occurrence of episodic desaturations in accordance with the increased REM phase during late postoperative nights [149] (Figure 18.9). Currently, the
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Figure 18.8 Changes in sleep architecture determined by pre- and postoperative full polysomnograms in a patient who had cholecystectomy under general anesthesia are presented. Compared with sleep pattern during the preoperative night, frequent arousal with lack of slow-wave sleep (SWS) and rapid eye movement sleep (REM) was observed during the operative night. Notably, significant increase in duration of REM sleep was evident during the third postoperative night (Data from Ref. 142).
REM rebound following REM suppression is considered a key pathogenic factor for development of late postoperative nocturnal hypoxemia. Anesthesia Technique and Postoperative Sleep Disturbance
Postoperative sleep disturbance could result from a variety of factors relating to anesthesia and surgical procedures, in addition to environmental factors such as noise, nursing procedures and light. Moote and Knill examined sleep before and after isoflurane anesthesia without surgery in healthy volunteers [182]. They found significant reduction of slow-wave sleep without influencing the proportion of REM sleep on the first postanesthetic night. REM suppression, which is a requirement for
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Figure 18.9 Recordings of arterial oxygen saturation (SpO2) and heart rate (HR) obtained in a 36-year-old woman on the second night after gastric resection are presented. Note episodic desaturation with marked heart rate variability during REM sleep (Data from Ref. 149).
development of the subsequent REM rebound, was not observed. Furthermore, Kavey and Ahshuler reported occurrence of REM rebound following REM suppression in patients who had herniorrhaphy with regional anesthesia [180]. Oxygenation during the second and third postoperative nights did not differ between patients receiving propofol anesthesia and those receiving isoflurane anesthesia, whereas better oxygenation during the first postoperative night was observed in patients receiving propofol [183]. Accordingly, anesthesia alone is unlikely to play a major role in the rebound increase in REM sleep during late postoperative nights. Opioids and Postoperative Sleep Disturbance
The contribution to late postoperative hypoxemia of opioids used for postoperative analgesia is controversial. Although only preliminary data published as an abstract, Moote et al. reported that morphine 0.2 mg/kg administered intramuscularly in healthy volunteers without pain caused marked sleep disturbance with significant reduction in both slow-wave sleep and REM sleep in addition to an increase in nocturnal awakening episodes, while a smaller dose (0.1 mg/kg) led to suppression of only slow-wave sleep [184]. Recently, this opioid-induced REM suppression was found to be mediated through m receptor subtypes in the medial pontine reticular formation in a cat preparation [185]. Despite the pharmacological effect
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of opioids, results of studies evaluating sleep on the second postoperative night indicate either a weak or no significant correlation between opioid dose and frequency of episodic desaturations [186,187]. Cronin et al. performed polysomnography on the preoperative and first three postoperative nights in patients undergoing low abdominal surgeries [187]. Severity and amount of REM rebound on the third postoperative night following abolishment of REM sleep on the first postoperative night did not differ between patients receiving epidural analgesia with fentanyl and those with bupivacaine alone, but the low plasma concentration of the opioid when administered into the epidural space might explain these negative observations. Accordingly, in our opinion, opioid administration is unlikely to be a key pathogenic factor for development of late postoperative hypoxemia, although postoperative use of a higher dose of opioid could be a contributing factor in selected patients. Surgical Stress and Postoperative Sleep Disturbance
Rosenberg et al. compared postoperative nocturnal oxygenation between patients with minor surgery (middle ear surgery) and those with major abdominal surgery [186]. Typical episodic hypoxemia was only observed in patients after major abdominal surgery. Thus, surgical stress is likely to be a major contributor to the REM rebound during late postoperative nights, a speculation supported by the fact that physical stress in nonsurgical patients with acute ischemic stroke [188] and acute myocardial infarction [189] produced similar sleep disturbances as those seen in postoperative patients. A variety of endocrine-metabolic and inflammatory mediators, which usually increase after surgery, were reported to influence sleep–wake regulation, mostly producing either sleep disturbance or REM suppression. For example, catecholamines appeared to play a role in maintenance of wakefulness [190], and administration of cortisol caused REM suppression with increasing NREM sleep in healthy volunteers [191]. Corticotrophin-releasing hormone decreased NREM sleep with increasing wakefulness in rabbits and rats [192,193]. While cytokines such as interleukin-1, interleukin-2 and interleukin-15 are somnogenic, REM sleep was suppressed in association with an increase in NREM sleep [194,195]. REM suppression was also reported after administration of endotoxin [196,197] and tumor necrosis factor [198]. Bauer et al. found significant REM suppression in patients with major depression during the first night following administration of endotoxin, which caused induction of cytokines and fever [197]. Although mood significantly improved during the next day, a rebound of REM sleep was observed during the second night after endotoxin administration and mood worsened again afterwards. Fever itself has significant influences on the sleep–wake regulation. Galland et al. demonstrated profound effects of artificially produced mild hyperthermia
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on REM sleep regulation and respiratory control [199]. REM sleep significantly decreased during hyperthermia and significantly increased during recovery from the hyperthermia. Apneas were commonly observed in association with the rebound REM during recovery from hyperthermia. C. Mechanisms of Late Postoperative Episodic Hypoxemia During Sleep
While mild hypoxemia physiologically develops during REM sleep, severe nocturnal hypoxemia observed after major surgery would not occur solely due to postoperative REM rebound. Either impairment of oxygenation and/ or emergence of or worsening of disordered breathing, possibly caused by surgical stress, need to be concomitantly present during the rebound REM period for deterioration of nocturnal oxygenation. Figure 18.10 illustrates hypothetical mechanisms for development of late postoperative episodic hypoxemia.
Surgical stress
Diaphragm dysfunction
Sleep deprivation
REM rebound
Reduction of FRC
Reduction of arousal response
Reduction of UA muscle activity
Increased UA collapsibility
Constant hypoxemia
Episodic hypoxemia due to UA obstruction
Figure 18.10 Hypothetical mechanisms of late postoperative episodic hypoxemia are illustrated assuming the nature of the postoperative sleep disordered breathing is obstructive. Rebound increase in REM sleep and FRC reduction after major abdominal surgery are the keys for development of the episodic desaturation due to upper airway (UA) obstruction.
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Impairment of Oxygenation due to Reduction of FRC
As mentioned above, the most significant change in the respiratory system, particularly after major abdominal surgery, is reduction of FRC, resulting in constant hypoxemia. Daytime blood gas analysis revealed lowest arterial oxygen tension on the second or third day after abdominal [200,201], thoracic [202], thoracoabdominal [203], and hip surgery [204]. Lung volume is a significant determinant for the rate of SaO2 reduction during apnea in addition to the initial SaO2 level. Accordingly, even a short period of apnea at lower FRC and SaO2 levels could result in severe hypoxemia postoperatively, while it would not cause hypoxemia preoperatively with normal starting FRC and SaO2. The reduced FRC would further decrease during the rebound REM period; Hudgel and Devadatta reported a decrease in FRC by 200–300 ml during REM sleep in normal humans [205]. Furthermore, upper airway resistance is reported to increase in association with reduction of lung volume [206], and therefore, the FRC reduction may itself be a predisposing factor for upper airway obstruction during the late postoperative period. Emergence or Worsening of Upper Airway Obstruction due to Increased Upper Airway Collapsibility
Assuming that the nature of disordered breathing causing episodic hypoxemia during late postoperative nights is obstructive, a variety of factors described below increase upper airway collapsibility. This could result in upper airway obstruction even in surgical patients without preoperative SDB or could lead to a worsening of upper airway obstruction in patients with preoperative SDB. First, sleep fragmentation caused by surgical stress could decrease the activity of pharyngeal airway dilating muscles, increasing collapsibility of the pharyngeal airway as demonstrated by Leiter et al. [207]. Furthermore, severe hypoxemia could be caused by prolonged pharyngeal obstruction, since depression of arousal in response to hypoxic, hypercapnic and laryngeal irritant stimuli has been demonstrated in dogs after sleep deprivation [208]. Second, increased time spent in the supine position postoperatively would be another predisposing factor for increasing pharyngeal airway collapsibility, although Rosenberg-Adamsen et al. failed to confirm this possibility in patients on the second postoperative night after major abdominal surgery [209]. D. Treatment Strategy for Late Postoperative Hypoxemia
An optimal treatment strategy for late postoperative hypoxemia has yet to be established, primarily because the nature of the disordered breathing causing the hypoxemia is still unclear. In fact, treatment may not be necessary for all surgical patients. We assume that patients with risk factors
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for development of late postoperative hypoxemia may require optimal management, particularly when these patients may develop postoperative organ dysfunction. Among the risk factors for episodic hypoxemia, preoperative SDB should be evaluated by polysomnography or nocturnal oximetry when suspected, based on clinical symptoms and body habitus [210]. Oxygen therapy for the episodic hypoxemia is controversial as is oxygen therapy for OSA [211,212]. While studies uniformly show improvement of mean SaO2 level by oxygen administration [147,213], Rosenberg et al. found no significant reduction of the number and duration of hypoxemia episodes with the therapy [213]. If episodic hypoxemia during the late postoperative period is primarily due to obstructive disordered breathing during sleep, nasal CPAP may be a rational and effective treatment option. While usefulness of nasal CPAP for management of OSA after surgery is widely accepted [214,215], no systematic assessment of nasal CPAP therapy for late postoperative hypoxemia has been performed. References 1.
2.
3. 4.
5.
6.
7.
8.
9.
Hines, R., Barash, P.G., Watrous, G. and O’Connor, T., Complications occurring in the postanesthesia care unit: a survey, Anesth. Analg. 74, 503–509, 1992. Asai, T., Koga, K. and Vaughan, R.S., Respiratory complications associated with tracheal intubation and extubation, Br. J. Anaesth. 80, 767– 775, 1998. Flemons, W.W., Obstructive sleep apnea, N. Engl. J. Med. 347, 498–504, 2002. Young, T., Palta, M., Dempsey, J., Skatrud, J., Weber, S. and Badr, S., The occurrence of sleep-disordered breathing among middle-aged adults, N. Engl. J. Med. 328, 1230–1235, 1993. Gupta, R.M., Parvizi, J., Hanssen, A.D. and Gay, P.C., Postoperative complications in patients with obstructive sleep apnea syndrome undergoing hip or knee replacement: a case–control study, Mayo Clin. Proc. 76, 897–905, 2001. Esclamado, R.M., Glenn, M.G., McCulloch, T.M. and Cummings, C.W., Perioperative complications and risk factors in the surgical treatment of obstructive sleep apnea syndrome, Laryngoscope 99, 1125–1129, 1989. Isono, S., Upper airway muscle function during sleep, in Sleep and Breathing in Children: A Developmental Approach, Loughlin, G.M., Marcus, C.L. and Carroll, J.L., eds., New York, Marcel Dekker, pp. 261–291, 2000. Kuna, S. and Remmers, J.E., Anatomy and physiology of upper airway obstruction, in Principles and Practice of Sleep Medicine, 3rd ed., Kryger, M.H., Roth, T. and Dement, W.C., eds., Philadelphia, WB Saunders, pp. 840–858, 2000. Remmers, J.E., DeGroot, W.J., Sauerland, E.K. and Anch, A.M., Pathogenesis of upper airway occlusion during sleep, J. Appl. Physiol. 44, 931–938, 1978.
Recovery from Anesthesia 10. 11.
12. 13.
14.
15.
16.
17.
18.
19.
20. 21.
22.
23. 24.
25.
26.
765
Brouillette, R.T. and Thach, B.T., A neuromuscular mechanism maintaining extrathoracic airway patency, J. Appl. Physiol. 46, 772–777, 1979. Van Lunteren, E., Van de Graaff, W.B., Parker, D.M., Mitra, J., Haxhiu, M.A., Strohl, K.P. and Cherniack, N.S., Nasal and laryngeal reflex responses to negative upper airway pressure, J. Appl. Physiol. 56, 746–752, 1984. Kuna, S.T., Inhibition of inspiratory upper airway motoneuron activity by phasic volume feedback, J. Appl. Physiol. 60, 1373–1379, 1986. Horner, R.L., Innes, J.A., Holden, H.B. and Guz, A., Afferent pathway(s) for pharyngeal dilator reflex to negative pressure in man: a study using upper airway anesthesia, J. Physiol. 436, 31–44, 1991. Parisi, R.A., Santiago, T.V. and Edelman, N.H., Genioglossal and diaphragmatic EMG responses to hypoxia during sleep, Am. Rev. Respir. Dis. 138, 610–616, 1988. Parisi, R.A., Neubauer, J.A., Frank, M.M., Edelman, N.H. and Santiago, T.V., Correlation between genioglossal and diaphragmatic responses to hypercapnia during sleep, Am. Rev. Respir. Dis. 135, 378–382, 1987. Wheatley, J.R., Tangel, D.J., Mezzanotte, W.S. and White, D.P., Influence of sleep on response to negative airway pressure of tensor palatini muscle and retropalatal airway, J. Appl. Physiol. 75, 2117–2124, 1993. Tangel, D.J., Mezzanotte, W.S. and White, D.P., Influence of sleep on tensor palatini EMG and upper airway resistance in normal men, J. Appl. Physiol. 70, 2574–2581, 1991. Isono, S., Remmers, J.E., Tanaka, A., Sho, Y., Sato, J. and Nishino, T., Anatomy of pharynx in patients with obstructive sleep apnea and normal subjects, J. Appl. Physiol. 82, 1319–1326, 1997. Isono, S., Tanaka, A., Sho, Y., Konno, A. and Nishino, T., Advancement of the mandible improves velopharyngeal airway patency, J. Appl. Physiol. 79, 2132–2138, 1995. McEvoy, R.D., Sharp, D.J. and Thornton, A.T., The effects of posture on obstructive sleep apnea, Am. Rev. Respir. Dis. 133, 662–666, 1986. Isono, S., Tanaka, A. and Nishino, T., Lateral position decreases collapsibility of the passive pharynx in patients with obstructive sleep apnea, Anesthesiology 97, 780–785, 2002. Ishikawa, T., Isono, S., Aiba, J., Tanaka, A. and Nishino, T., Prone position increases collapsibility of the passive pharynx in infants and small children, Am. J. Respir. Crit. Care Med. 166, 760–764, 2002. Van de Graaff, W.B., Thoracic influence on upper airway patency, J. Appl. Physiol. 65, 2124–2134, 1988. Thut, D.C., Schwartz, A.R., Roach, D., Wise, R.A., Permutt, S. and Smith, P.L., Tracheal and neck position influence upper airway airflow dynamics by altering airway length, J. Appl. Physiol. 75, 2084–2090, 1993. Cala, S.J., Sliwinski, P., Cosio, M.G. and Kimoff, R.J., Effects of topical upper airway anesthesia on apnea duration through the night in obstructive sleep apnea, J. Appl. Physiol. 81, 2618–2626, 1996. Berry, R.B., Kouchi, K.G., Bower, J.L. and Light, R.W., Effects of upper airway anesthesia on obstructive sleep apnea, Am. J. Respir. Crit. Care Med. 151, 1857–1861, 1995.
766 27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39. 40.
41.
42.
Isono and Rosenberg Nishino, T., Shirahata, M., Yonezawa, T. and Honda, Y., Comparison of changes in the hypoglossal and the phrenic nerve activity in response to increasing depth of anesthesia in cats, Anesthesiology 60, 19–24, 1984. Catley, D.M., Thornton, C., Jordan, C., Lehane, J.R., Royston, D. and Jones, J.G., Pronounced, episodic oxygen desaturation in the postoperative period: its association with ventilatory pattern and analgesic regimen, Anesthesiology 63, 20–28, 1985. Parr, S.M., Robinson, B.J., Glover, P.W. and Galletly, D.C., Level of consciousness on arrival in the recovery room and the development of early respiratory morbidity, Anaesth. Intens. Care 19, 369–372, 1991. Bowes, G., Townsend, E.R., Bromley, S.M., Kozar, L.F. and Phillipson, E.A., Role of the carotid body and of afferent vagal stimuli in the arousal response to airway occlusion in sleeping dogs, Am. Rev. Respir. Dis. 123, 644–647, 1981. Knill, R.L. and Gelb, A.W., Ventilatory responses to hypoxia and hypercapnia during halothane sedation and anesthesia in man, Anesthesiology 49, 244–251, 1978. Wymore, M.L. and Eisele, J.H., Differential effects of d-tubocurarine on inspiratory muscles and two peripheral muscle groups in anesthetized man, Anesthesiology 48, 360–362, 1978. Donati, F., Antzaka, C. and Bevan, D.R., Potency of pancuronium at the diaphragm and the adductor pollicis muscle in humans, Anesthesiology 65, 1–5, 1986. Isono, S., Kochi, T., Ide, T., Sugimori, K., Mizuguchi, T. and Nishino, T., Differential effects of vecuronium on diaphragm and geniohyoid muscle in anaesthetized dogs, Br. J. Anaesth. 68, 239–243, 1992. D’Honneur, G., Lofaso, F., Drummond, G.B., Rimaniol, J.M., Aubineau, J.V., Harf, A. and Duvaldestin, P., Susceptibility to upper airway obstruction during partial neuromuscular block, Anesthesiology 88, 371–378, 1998. Eriksson, L.I., Sato, M. and Severinghaus, J.W., Effect of a vecuroniuminduced partial neuromuscular block on hypoxic ventilatory response, Anesthesiology 78, 693–699, 1993. Igarashi, A., Amagasa, S., Horikawa, H. and Shirahata, M., Vecuronium directly inhibits hypoxic neurotransmission of the rat carotid body, Anesth. Analg. 94, 117–122, 2002. Riley, R.W., Powell, N.B., Guilleminault, C., Pelayo, R., Troell, R.J. and Li, K.K., Obstructive sleep apnea surgery: risk management and complications, Otolaryngol. Head Neck Surg. 117, 648–652, 1997. Fairbanks, D.N.F., Uvulopalatopharyngoplasty complications and avoidance strategies, Otolaryngol. Head Neck Surg. 102, 239–245, 1990. Sagi, H.C., Beutler, W., Carroll, E. and Connolly, P.J., Airway complications associated with surgery on the anterior cervical spine, Spine 27, 949– 953, 2002. Van der Touw, T., Crawford, A.B. and Wheatley, J.R., Effects of a synthetic lung surfactant on pharyngeal patency in awake human subjects, J. Appl. Physiol. 82, 78–85, 1997. Olson, L.G. and Strohl, K.P., Airway secretions influence upper airway patency in the rabbit, Am. Rev. Respir. Dis. 137, 1379–1381, 1988.
Recovery from Anesthesia 43.
44. 45.
46.
47.
48.
49.
50.
51.
52.
53. 54.
55.
56. 57. 58.
767
Boudewyns, A., Punjabi, N., Van de Heyning, P.H., De Backer, W.A., O’Donnell. C.P., Schneider, H., Smith, P.L. and Schwartz, A.R., Abbreviated method for assessing upper airway function in obstructive sleep apnea, Chest 118, 1031–1041, 2000. Safar, P., Escarraga, L.A. and Chang, F., Upper airway obstruction in the unconscious patient, J. Appl. Physiol. 14, 760–764, 1959. Shorten, G.D., Armstrong, D.C., Roy, W.I. and Brown, L., Assessment of the effect of head and neck position on upper airway anatomy in sedated paediatric patients using magnetic resonance imaging, Paediatr. Anaesth. 5, 243–248, 1995. Sivarajan, M. and Joy, J.V., Effects of general anesthesia and paralysis on upper airway changes due to head position in humans, Anesthesiology 85, 787–793, 1996. Olsson, G.L. and Hallen, B., Laryngospasm during anaesthesia. A computeraided incidence study in 136,929 patients, Acta Anaesthesiol. Scand. 28, 567–575, 1984. Tait, A.R., Malviya, S., Voepel-Lewis, T., Munro, H.M., Seiwert, M. and Pandit, U.A., Risk factors for perioperative adverse respiratory events in children with upper respiratory tract infections, Anesthesiology 95, 299–306, 2001. Pounder, D.R., Blackstock, D. and Steward, D.J., Tracheal extubation in children: halothane versus isoflurane, anesthetized versus awake, Anesthesiology 74, 653–655, 1991. Schreiner, M.S., O’Hara, I., Markakis, D.A. and Politis, G.D., Do children who experience laryngospasm have an increased risk of upper respiratory tract infection? Anesthesiology 85, 475–480, 1996. Patel, R.I., Hannallah, R.S., Norden, J., Casey, W.F. and Verghese, S.T., Emergence airway complications in children: a comparison of tracheal extubation in awake and deeply anesthetized patients, Anesth. Analg. 73, 266–270, 1991. Halow, K.D. and Ford, E.G., Pulmonary edema following post-operative laryngospasm: a case report and review of the literature, Am. Surg. 59: 443–447, 1993. Baraka, A., Intravenous lidocaine controls extubation laryngospasm in children, Anesth. Analg. 57, 506–507, 1978. Leicht, P., Wisborg, T. and Chraemmer-Jorgensen, B., Does intravenous lidocaine prevent laryngospasm after extubation in children? Anesth. Analg. 64, 1193–1196, 1985. Afshan, G., Chohan, U., Qamar-Ul-Hoda, M. and Kamal, R.S., Is there a role of a small dose of propofol in the treatment of laryngeal spasm? Paediatr. Anaesth. 12, 625–628, 2002. Miller, K.A., Harkin, C.P. and Bailey, P.L., Postoperative tracheal extubation, Anesth. Analg. 80, 149–172, 1995. Bishop, M.J., Weymuller, E.A., Jr. and Fink, B.R., Laryngeal effects of prolonged intubation, Anesth. Analg. 63, 335–342, 1984. Tanaka, A., Isono, S., Sato, J. and Nishino, T., Effects of minor surgery and endotracheal intubation on postoperative breathing patterns in patients anaesthetized with isoflurane or sevoflurane, Br. J. Anaesth. 87, 706–710, 2001.
768 59.
60.
61.
62. 63. 64. 65.
66. 67.
68. 69.
70. 71.
72.
73. 74.
75.
Isono and Rosenberg Tanaka, A., Isono, S., Ishikawa, T., Sato, J. and Nishino, T., Laryngeal resistance before and after minor surgery: endotracheal tube versus laryngeal mask airway, Anesthesiology 99, 252–258, 2003. Weymuller, E.A., Jr., Bishop, M.J., Fink, B.R., Hibbard, A.W. and Spelman, F.A., Quantification of intralaryngeal pressure exerted by endotracheal tubes, Ann. Otol. Rhinol. Laryngol. 92, 444–447, 1983. Litman, R.S., Weissend, E.E., Shibata, D. and Westesson, P.L., Developmental changes of laryngeal dimensions in unparalyzed, sedated children, Anesthesiology 98, 41–45, 2003. Koka, B.V., Jeon, I.S., Andre, J.M., MacKay, I. and Smith, R.M., Postintubation croup in children, Anesth. Analg. 56, 501–505, 1977. Fisher, M.M. and Raper, R.F., The ‘cuff-leak’ test for extubation, Anaesthesia 47, 10–12, 1992. Miller, R.L. and Cole, R.P., Association between reduced cuff leak volume and postextubation stridor, Chest 110, 1035–1040, 1996. Davis, P.G. and Henderson-Smart, D.J., Intravenous dexamethasone for extubation of newborn infants, Cochrane Database Syst. Rev. 4, CD000308, 2001. Davies, M.W. and Davis, P.G., Nebulized racemic epinephrine for extubation of newborn infants, Cochrane Database Syst. Rev. 1, CD000506, 2002. Darmon, J.Y., Rauss, A., Dreyfuss, D., Bleichner, G., Elkharrat, D., Schlemmer, B., Tenaillon, A., Brun-Buisson, C. and Huet, Y., Evaluation of risk factors for laryngeal edema after tracheal extubation in adults and its prevention by dexamethasone. A placebo-controlled, double-blind, multicenter study, Anesthesiology 77, 245–251, 1992. Efferen, L.S. and Elsakr, A., Post-extubation stridor: risk factors and outcome, J. Assoc. Acad. Minor. Phys. 9, 65–68, 1998. Roberts, K.W., Crnkovic, A. and Steiniger, J.R., Post-anesthesia paradoxical vocal cord motion successfully treated with midazolam, Anesthesiology 89, 517– 519, 1998. Hammer, G., Schwinn, D. and Wollman, H., Postoperative complications due to paradoxical vocal cord motion, Anesthesiology 66, 686–687, 1987. Kakitsuba, N., Sadaoka, T., Kanai, R., Fujiwara, Y. and Takahashi, H., Peculiar snoring in patients with multiple system atrophy: its sound source, acoustic characteristics, and diagnostic significance, Ann. Otol. Rhinol. Laryngol. 106, 380–384, 1997. Isono, S., Shiba, K., Yamaguchi, M., Tanaka, A., Hattori, T., Konno, A. and Nishino, T., Pathogenesis of laryngeal narrowing in patients with multiple system atrophy, J. Physiol. 536, 237–249, 2001. Tomlin, P.J., Howarth, F.H. and Robinson, J.S., Postoperative atelectasis and laryngeal incompetence, Lancet 1, 1402–1405, 1968. Burgess, G.E., III, Cooper, J.R., Jr., Marino, R.J., Peuler, M.J. and Warriner, R.A., III, Laryngeal competence after tracheal extubation, Anesthesiology 51, 73–77, 1979. Stanley, G.D., Bastianpillai, B.A., Mulcahy, K. and Langton, J.A., Postoperative laryngeal competence. The laryngeal mask airway and tracheal tube compared, Anaesthesia 50, 985–986, 1995.
Recovery from Anesthesia 76.
77.
78.
79.
80.
81.
82.
83.
84. 85. 86.
87.
88.
89.
90.
769
Nishino, T., Takizawa, K., Yokokawa, N. and Hiraga, K., Depression of the swallowing reflex during sedation and/or relative analgesia produced by inhalation of 50% nitrous oxide in oxygen, Anesthesiology 67, 995–998, 1987. Nishino, T., Hiraga, K. and Yokokawa, N., Laryngeal and respiratory responses to tracheal irritation at different depths of enflurane anesthesia in humans, Anesthesiology 73, 46–51, 1990. Pavlin, E.G., Holle, R.H. and Schoene, R.B., Recovery of airway protection compared with ventilation in humans after paralysis with curare, Anesthesiology 70, 381–385, 1989. Isono, S., Ide, T., Kochi, T., Mizuguchi, T. and Nishino, T., Effects of partial paralysis on the swallowing reflex in conscious humans, Anesthesiology 75, 980– 984, 1991. Eriksson, L.I., Sundman, E., Olsson, R., Nilsson, L., Witt, H., Ekberg, O. and Kuylenstierna, R., Functional assessment of the pharynx at rest and during swallowing in partially paralyzed humans: simultaneous videomanometry and mechanomyography of awake human volunteers, Anesthesiology 87, 1035–1043, 1997. Hasegawa, R. and Nishino, T., Temporal changes in airway protective reflexes elicited by an endotracheal tube in surgical patients anaesthetized with sevoflurane, Eur. J. Anaesthesiol. 16, 98–102, 1999. Davies, R.O., Edwards, M.W., Jr. and Lahiri, S., Halothane depresses the response of carotid body chemoreceptors to hypoxia and hypercapnia in the cat, Anesthesiology 57, 153–159, 1982. Ide, T., Sakurai, Y., Aono, M. and Nishino, T., Contribution of peripheral chemoreception to the depression of the hypoxic ventilatory response during halothane anesthesia in cats, Anesthesiology 90, 1084–1091, 1999. Knill, R.L. and Gelb, A.W., Peripheral chemoreceptors during anesthesia: are the watchdogs sleeping? Anesthesiology 57, 151–152, 1982. Baraka, A., Postoperative respiratory depression following excision of carotid body tumours, Can. J. Anaesth. 41, 306–309, 1994. Weil, J.V., McCullough, R.E., Kline, J.S. and Sodal, I.E., Diminished ventilatory response to hypoxia and hypercapnia after morphine in normal man, N. Engl. J. Med. 292, 1103–1106, 1975. Bailey, P.L., Lu, J.K., Pace, N.L., Orr, J.A., White, J.L., Hamber, E.A., Slawson, M.H., Crouch, D.J. and Rollins, D.E., Effects of intrathecal morphine on the ventilatory response to hypoxia, N. Engl. J. Med. 343, 1228–1234, 2000. McQueen, D.S. and Ribeiro, J.A., Inhibitory actions of methionine-enkephalin and morphine on the cat carotid chemoreceptors, Br. J. Pharmacol. 71, 297–305, 1980. Becker, L.D., Paulson, B.A., Miller, R.D., Severinghaus, J.W. and Eger, E.I., II, Biphasic respiratory depression after fentanyl–droperidol or fentanyl alone used to supplement nitrous oxide anesthesia, Anesthesiology 44, 291– 296, 1976. Mahla, M.E., White, S.E. and Moneta, M.D., Delayed respiratory depression after alfentanil, Anesthesiology 69, 593–595, 1988.
770
Isono and Rosenberg
91.
Stoeckel, H., Hengstmann, J.H. and Schuttler, J., Pharmacokinetics of fentanyl as a possible explanation for recurrence of respiratory depression, Br. J. Anaesth. 51, 741–745, 1979. Smedstad, K.G. and Rigg, J.R., Control of breathing after fentanyl and Innovar anaesthesia, Br. J. Anaesth. 54, 599–604, 1982. Lam, A.M., Clement, J.L. and Knill, R.L., Surgical stimulation does not enhance ventilatory chemoreflexes during enflurane anaesthesia in man, Can. Anaesth. Soc. J. 27, 22–28, 1980. Nishino, T. and Kochi, T., Effects of surgical stimulation on the apnoeic thresholds for carbon dioxide during anaesthesia with sevoflurane, Br. J. Anaesth. 73, 583–586, 1994. Sarton, E., Dahan, A., Teppema, L., van den Elsen, M., Olofsen, E., Berkenbosch, A. and van Kleef, J., Acute pain and central nervous system arousal do not restore impaired hypoxic ventilatory response during sevoflurane sedation, Anesthesiology 85, 295–303, 1996. Borgbjerg, F.M., Nielsen, K. and Franks, J., Experimental pain stimulates respiration and attenuates morphine-induced respiratory depression: a controlled study in human volunteers, Pain 64, 123–128, 1996. Hedenstierna, G., Strandberg, A., Brismar, B., Lundquist, H., Svensson, L. and Tokics, L., Functional residual capacity, thoracoabdominal dimensions, and central blood volume during general anesthesia with muscle paralysis and mechanical ventilation, Anesthesiology 62, 247–254, 1985. Froese, A.B. and Bryan, A.C., Effects of anesthesia and paralysis on diaphragmatic mechanics in man, Anesthesiology 41, 242–255, 1974. Muller, N., Volgyesi, G., Becker, L., Bryan, M.H. and Bryan, A.C., Diaphragmatic muscle tone, J. Appl. Physiol. 47, 279–284, 1979. Ali, J., Weisel, R.D., Layug, A.B., Kripke, B.J. and Hechtman, H.B., Consequences of postoperative alterations in respiratory mechanics, Am. J. Surg. 128, 376–382, 1974. Meyers, J.R., Lembeck, L., O’Kane, H. and Baue, A.E., Changes in functional residual capacity of the lung after operation, Arch. Surg. 110, 576–583, 1975. Craig, D.B., Postoperative recovery of pulmonary function, Anesth. Analg. 60, 46–52, 1981. Hedenstierna, G. and Lofstrom, J., Effect of anaesthesia on respiratory function after major lower extremity surgery. A comparison between bupivacaine spinal analgesia with low-dose morphine and general anaesthesia, Acta Anaesthesiol. Scand. 29, 55–60, 1985. Putensen-Himmer, G., Putensen, C., Lammer, H., Lingnau, W., Aigner, F. and Benzer, H., Comparison of postoperative respiratory function after laparoscopy or open laparotomy for cholecystectomy, Anesthesiology 77, 675– 680, 1992. Eichenberger, A., Proietti, S., Wicky, S., Frascarolo, P., Suter, M., Spahn, D.R. and Magnusson, L., Morbid obesity and postoperative pulmonary atelectasis: an underestimated problem, Anesth. Analg. 95, 1788–1792, 2002. Ford, G.T., Whitelaw, W.A., Rosenal, T.W., Cruse, P.J. and Guenter, C.A., Diaphragm function after upper abdominal surgery in humans, Am. Rev. Respir. Dis. 127, 431–436, 1983.
92. 93.
94.
95.
96.
97.
98. 99. 100.
101. 102. 103.
104.
105.
106.
Recovery from Anesthesia
771
107. Dureuil, B., Viires, N., Cantineau, J.P., Aubier, M. and Desmonts, J.M., Diaphragmatic contractility after upper abdominal surgery, J. Appl. Physiol. 61, 1775–1780, 1986. 108. Oyer, L.M., Knuth, S.L., Ward, D.K. and Bartlett, D., Jr., Reflex inhibition of crural diaphragmatic activity by esophageal distention in cats, Respir. Physiol. 77, 195–202, 1989. 109. Prabhakar, N.R., Marek, W. and Loeschcke, H.H., Altered breathing pattern elicited by stimulation of abdominal visceral afferents, J. Appl. Physiol. 58, 1755–1760, 1985. 110. Ford, G.T., Grant, D.A., Rideout, K.S., Davison, J.S. and Whitelaw, W.A., Inhibition of breathing associated with gallbladder stimulation in dogs, J. Appl. Physiol. 65, 72–79, 1988. 111. Spence, A.A. and Smith, G., Postoperative analgesia and lung function: a comparison of morphine with extradural block. Br. J. Anaesth. 43, 144–148, 1971. 112. Wahba, W.M., Don, H.F. and Craig, D.B., Post-operative epidural analgesia: effects on lung volumes, Can. Anaesth. Soc. J. 22, 519–527, 1975. 113. Manikian, B., Cantineau, J.P., Bertrand, M., Kieffer, E., Sartene, R. and Viars, P., Improvement of diaphragmatic function by a thoracic extradural block after upper abdominal surgery, Anesthesiology 68, 379–386, 1988. 114. Jayr, C., Thomas, H., Rey, A., Farhat, F., Lasser, P. and Bourgain, J.L., Postoperative pulmonary complications. Epidural analgesia using bupivacaine and opioids versus parenteral opioids, Anesthesiology 78, 666– 676, 1993. 115. Gamsu, G., Singer, M.M., Vincent, H.H., Berry, S. and Nadel, J.A., Postoperative impairment of mucous transport in the lung, Am. Rev. Respir. Dis. 114, 673–679, 1976. 116. Forbes, A.R., Halothane depresses mucociliary flow in the trachea, Anesthesiology 45, 59–63, 1976. 117. Raphael, J.H., Selwyn, D.A., Mottram, S.D., Langton, J.A. and O’Callaghan, C., Effects of 3 MAC of halothane, enflurane and isoflurane on cilia beat frequency of human nasal epithelium in vitro, Br. J. Anaesth. 76, 116–121, 1996. 118. Raphael, J.H., Strupish, J., Selwyn, D.A., Hann, H.C. and Langton, J.A., Recovery of respiratory ciliary function after depression by inhalation anaesthetic agents: an in vitro study using nasal turbinate explants, Br. J. Anaesth. 76, 854–859, 1996. 119. Hann, H.C., Hall, A.P., Raphael, J.H. and Langton, J.A., An investigation into the effects of midazolam and propofol on human respiratory cilia beat frequency in vitro. Intens. Care Med. 24, 791–794, 1998. 120. Konrad, F., Schreiber, T., Grunert, A., Clausen, M. and Ahnefeld, F.W., Measurement of mucociliary transport velocity in ventilated patients. Shortterm effect of general anesthesia on mucociliary transport, Chest 102, 1377–1383, 1992. 121. Forbes, A.R. and Gamsu, G., Depression of lung mucociliary clearance by thiopental and halothane, Anesth. Analg. 58, 387–389, 1979.
772
Isono and Rosenberg
122. Branson, R.D., Campbell, R.S., Davis, K. and Porembka, D.T., Anaesthesia circuits, humidity output, and mucociliary structure and function, Anaesth. Intens. Care 26, 178–183, 1998. 123. Forbes, A.R. and Gamsu, G., Lung mucociliary clearance after anesthesia with spontaneous and controlled ventilation, Am. Rev. Respir. Dis. 120, 857–862, 1979. 124. Alexopoulos, C., Jansson, B. and Lindholm, C.E., Mucus transport and surface damage after endotracheal intubation and tracheostomy. An experimental study in pigs, Acta Anaesthesiol. Scand. 28, 68–76, 1984. 125. Konrad, F., Schiener, R., Marx, T. and Georgieff, M., Ultrastructure and mucociliary transport of bronchial respiratory epithelium in intubated patients, Intens. Care Med. 21, 482–489, 1995. 126. Puchelle, E., Zahm, J.M. and Bertrand, A., Influence of age on bronchial mucociliary transport, Scand. J. Respir. Dis. 60, 307–313, 1979. 127. Konrad, F.X., Schreiber, T., Brecht-Kraus, D. and Georgieff, M., Bronchial mucus transport in chronic smokers and nonsmokers during general anesthesia, J. Clin. Anesth. 5, 375–380, 1993. 128. Brooks-Brunn, J.A., Predictors of postoperative pulmonary complications following abdominal surgery, Chest 111, 564–571, 1997. 129. Lawrence, V.A., Dhanda, R., Hilsenbeck, S.G. and Page, C.P., Risk of pulmonary complications after elective abdominal surgery, Chest 110, 744–750, 1996. 130. Shackford, S.R., Virgilio, R.W. and Peters, R.M., Early extubation versus prophylactic ventilation in the high-risk patient: a comparison of postoperative management in the prevention of respiratory complications, Anesth. Analg. 60, 76–80, 1981. 131. Hedstrand, U., Liw, M., Rooth, G. and Ogren, C.H., Effect of respiratory physiotherapy on arterial oxygen tension, Acta Anaesthesiol. Scand. 22, 349–352, 1978. 132. Chumillas, S., Ponce, J.L., Delgado, F., Viciano, V. and Mateu, M., Prevention of postoperative pulmonary complications through respiratory rehabilitation: a controlled clinical study, Arch. Phys. Med. Rehabil. 79, 5–9, 1998. 133. Gosselink, R., Schrever, K., Cops, P., Witvrouwen, H., De Leyn, P., Troosters, T., Lerut, A., Deneffe, G. and Decramer, M., Incentive spirometry does not enhance recovery after thoracic surgery, Crit. Care Med. 28, 679–683, 2000. 134. Ali, J., Serrette, C., Wood, L.D. and Anthonisen, N.R., Effect of postoperative intermittent positive pressure breathing on lung function, Chest 85, 192–196, 1984. 135. Denehy, L., Carroll, S., Ntoumenopoulos, G. and Jenkins, S., A randomized controlled trial comparing periodic mask CPAP with physiotherapy after abdominal surgery, Physiother. Res. Int. 6, 236–250, 2001. 136. Thomas, J.A. and McIntosh, J.M., Are incentive spirometry, intermittent positive pressure breathing, and deep breathing exercises effective in the prevention of postoperative pulmonary complications after upper abdominal surgery? A systematic overview and meta-analysis, Phys. Ther. 74, 3–10, 1994.
Recovery from Anesthesia
773
137. Jansen, J.E., Sorensen, A.I., Naesh, O., Erichsen, C.J. and Pedersen, A., Effect of doxapram on postoperative pulmonary complications after upper abdominal surgery in high-risk patients, Lancet 335, 936–938, 1990. 138. Dureuil, B., Desmonts, J.M., Mankikian, B. and Prokocimer, P., Effects of aminophylline on diaphragmatic dysfunction after upper abdominal surgery, Anesthesiology 62, 242–246, 1985. 139. Fujii, Y., Toyooka, H. and Amaha, K., Amrinone improves contractility of fatigued diaphragm in dogs, Can. J. Anaesth. 42, 80–86, 1995. 140. Uemura, A., Fujii, Y. and Toyooka, H., Inhaled olprinone improves contractility of fatigued canine diaphragm, Br. J. Anaesth. 88, 408–411, 2002. 141. Rosenberg, J., Dirkes, W.E. and Kehlet, H., Episodic arterial oxygen desaturation and heart rate variations following major abdominal surgery, Br. J. Anaesth. 63, 651–654, 1989. 142. Knill, R.L., Moote, C.A., Skinner, M.I. and Rose, E.A., Anesthesia with abdominal surgery leads to intense REM sleep during the first postoperative week, Anesthesiology 73, 52–61, 1990. 143. Douglas, N.J., Thomas, S. and Jan, M.A., Clinical value of polysomnography, Lancet 339, 347–350, 1992. 144. Gyulay, S., Olson, L.G., Hensley, M.J., King, M.T., Allen, K.M. and Saunders, N.A., A comparison of clinical assessment and home oximetry in the diagnosis of obstructive sleep apnea, Am. Rev. Respir. Dis. 147, 50–53, 1993. 145. Rosenberg, J., Rasmussen, G.I., Wojdemann, K.R., Kirkeby, L.T., Jorgensen, L.N. and Kehlet, H., Ventilatory pattern and associated episodic hypoxaemia in the late postoperative period in the general surgical ward, Anaesthesia 54, 323–328, 1999. 146. Rosenberg, J., Pedersen, M.H., Ullstad, T., von Jessen, F., Jepersen, N., Vinge, O., Rasmussen, J., Hjorne, F.P., Poulsen, N.J. and Kehlet, H., Incidence of arterial hypoxemia after laparotomy, Surg. Forum 43, 35–37, 1992. 147. Reeder, M.K., Goldman, M.D., Loh, L., Muir, A.D., Foex, P., Casey, K.R. and McKenzie, P.J., Postoperative hypoxaemia after major abdominal vascular surgery, Br. J. Anaesth. 68, 23–26, 1992. 148. Rosenberg, J., Rasmussen, V., von Jessen, F., Ullstad, T. and Kehlet, H., Late postoperative episodic and constant hypoxaemia and associated ECG abnormalities, Br. J. Anaesth. 65, 684–691, 1990. 149. Rosenberg, J., Wildschiodtz, G., Pedersen, M.H., von Jessen, F. and Kehlet, H., Late postoperative nocturnal episodic hypoxaemia and associated sleep pattern, Br. J. Anaesth. 72, 145–150, 1994. 150. Rosenberg, J., Ullstad, T., Rasmussen, J., Hjorne, F.P., Poulsen, N.J. and Goldman, M.D., Time course of postoperative hypoxaemia, Eur. J. Surg. 160, 137–143, 1994. 151. Isono, S., Sha, M., Suzukawa, M., Sho, Y., Ohmura, A., Kudo, Y., Misawa, K., Inaba, S. and Nishino, T., Preoperative nocturnal desaturations as a risk factor for late postoperative nocturnal desaturations, Br. J. Anaesth. 80, 602–605, 1998.
774
Isono and Rosenberg
152. Gentil, B., Lienhart, A. and Fleury, B., Enhancement of postoperative desaturation in heavy snorers, Anesth. Analg. 81, 389–392, 1995. 153. Pateman, J.A. and Hanning, C.D., Postoperative myocardial infarction and episodic hypoxaemia, Br. J. Anaesth. 63, 648–650, 1989. 154. Tarhan, S., Moffitt, E.A., Taylor, W.F. and Giuliani, E.R., Myocardial infarction after general anesthesia, JAMA 220, 1451–1454, 1972. 155. Reeder, M.K., Muir, A.D., Foex, P., Goldman, M.D., Loh, L. and Smart, D., Postoperative myocardial ischaemia: temporal association with nocturnal hypoxaemia, Br. J. Anaesth. 67, 626–631, 1991. 156. Gill, N.P., Wright, B. and Reilly, C.S., Relationship between hypoxaemic and cardiac ischaemic events in the perioperative period, Br. J. Anaesth. 68, 471–473, 1992. 157. Galatius-Jensen, S., Hansen, J., Rasmussen, V., Bildsoe, J., Therboe, M. and Rosenberg, J., Nocturnal hypoxaemia after myocardial infarction: association with nocturnal myocardial ischaemia and arrhythmias, Br. Heart J. 72, 23–30, 1994. 158. Rosenberg-Adamsen, S., Lie, C., Bernhard, A., Kehlet, H. and Rosenberg, J., Effect of oxygen treatment on heart rate after abdominal surgery, Anesthesiology 90, 380–384, 1999. 159. Guilleminault, C., Connolly, S.J. and Winkle, R.A., Cardiac arrhythmia and conduction disturbances during sleep in 400 patients with sleep apnea syndrome, Am. J. Cardiol. 52, 490–494, 1983. 160. Rosenberg, J., Pedersen, M.H., Ramsing, T. and Kehlet, H., Circadian variation in unexpected postoperative death, Br. J. Surg. 79, 1300–1302, 1992. 161. Gogenur, I., Rosenberg-Adamsen, S., Lie, C., Rasmussen, V. and Rosenberg, J., Lack of circadian variation in the activity of the autonomic nervous system after major abdominal operations, Eur. J. Surg. 168, 242–246, 2002. 162. Franco, P., Szliwowski, H., Dramaix, M. and Kahn, A., Polysomnographic study of the autonomic nervous system in potential victims of sudden infant death syndrome, Clin. Auton. Res. 8, 243–249, 1998. 163. Aakerlund, L.P. and Rosenberg, J., Postoperative delirium: treatment with supplementary oxygen, Br. J. Anaesth. 72, 286–290, 1994. 164. Marcantonio, E.R., Goldman, L., Mangione, C.M., Ludwig, L.E., Muraca, B., Haslauer, C.M., Donaldson, M.C., Whittemore, A.D., Sugarbaker, D.J. and Poss, R., et al., A clinical prediction rule for delirium after elective noncardiac surgery, JAMA 271, 134–139, 1994. 165. Schor, J.D., Levkoff, S.E., Lipsitz, L.A., Reilly, C.H., Cleary, P.D., Rowe, J.W. and Evans, D.A., Risk factors for delirium in hospitalized elderly, JAMA 267, 827–831, 1992. 166. Dieckelmann, A., Haupts, M., Kaliwoda, A., Rembs, E., Haan, J. and Zumtobel, V., Acute postoperative psychosyndromes. A prospective study and multivariate analysis of risk factors, Chirurg. 60, 470–474, 1989. 167. Ghoneim, M.M., Hinrichs, J.V., O’Hara, M.W., Mehta, M.P., Pathak, D., Kumar, V. and Clark, C.R., Comparison of psychologic and cognitive functions after general or regional anesthesia, Anesthesiology 69, 507–515, 1988.
Recovery from Anesthesia
775
168. Williams-Russo, P., Urquhart, B.L., Sharrock, N.E. and Charlson, M.E., Post-operative delirium: predictors and prognosis in elderly orthopedic patients, J. Am. Geriatr. Soc. 40, 759–767, 1992. 169. Berggren, D., Gustafson, Y., Eriksson, B., Bucht, G., Hansson, L.I., Reiz, S. and Winblad, B., Postoperative confusion after anesthesia in elderly patients with femoral neck fractures, Anesth. Analg. 66, 497–504, 1987. 170. Gibson, G.E., Pulsinelli, W., Blass, J.P. and Duffy, T.E., Brain dysfunction in mild to moderate hypoxia, Am. J. Med. 70, 1247–1254, 1981. 171. Rosenberg, J. and Kehlet, H., Postoperative mental confusion — association with postoperative hypoxemia, Surgery 114, 76–81, 1993. 172. Krasheninnikoff, M., Ellitsgaard, N., Rude, C. and Moller, J.T., Hypoxaemia after osteosynthesis of hip fractures, Int. Orthop. 17, 27–29, 1993. 173. Condon, R.E. and Wittmann, D.H., Surgical infections, in Oxford Textbook of Surgery, Morris, P.J. and Malt, R.A., eds., New York, Oxford University Press, pp. 27–43, 1994. 174. Jonsson, K., Jensen, J.A., Goodson, W.H., III, Scheuenstuhl, H., West, J., Hopf, H.W. and Hunt, T.K., Tissue oxygenation, anemia, and perfusion in relation to wound healing in surgical patients, Ann. Surg. 214, 605–613, 1991. 175. Knighton, D.R., Fiegel, V.D., Halverson, T., Schneider, S., Brown, T. and Wells, C.L., Oxygen as an antibiotic. The effect of inspired oxygen on bacterial clearance, Arch. Surg. 125, 97–100, 1990. 176. Orgill, D. and Demling, R.H., Current concepts and approaches to wound healing, Crit. Care Med. 16, 899–908, 1988. 177. Rosenberg, J., Pedersen, U., Erichsen, C.J., Vibits, H., Moesgaard, F. and Kehlet, H., Effect of epidural blockade and oxygen therapy on changes in subcutaneous oxygen tension after abdominal surgery, J. Surg. Res. 56, 72–76, 1994. 178. Hartmann, M., Jonsson, K. and Zederfeldt, B., Effect of tissue perfusion and oxygenation on accumulation of collagen in healing wounds. Randomized study in patients after major abdominal operations, Eur. J. Surg. 158, 521–526, 1992. 179. Rosenberg, J., Ullstad, T., Larsen, P.N., Moesgaard, F. and Kehlet, H., Continuous assessment of oxygen saturation and subcutaneous oxygen tension after abdominal operations, Acta Chir. Scand. 156, 585–590, 1990. 180. Kavey, N.B. and Ahshuler, K.Z., Sleep in herniorrhaphy patients, Am. J. Surg. 138, 683–687, 1979. 181. Dickerson, L.W., Huang, A.H., Thurnher, M.M., Nearing, B.D. and Verrier, R.L., Relationship between coronary hemodynamic changes and the phasic events of rapid eye movement sleep, Sleep 16, 550–557, 1993. 182. Moote, C.A. and Knill, R.L., Isoflurane anesthesia causes a transient alteration in nocturnal sleep, Anesthesiology 69, 327–331, 1988. 183. Georgiou, L.G., Vourlioti, A.N., Kremastinou, F.I., Stefanou, P.S., Tsiotou, A.G. and Kokkinou, M.D., Influence of anesthetic technique on early postoperative hypoxemia, Acta Anaesthesiol. Scand. 40, 75–80, 1996. 184. Moote, C.A., Knill, R.L., Skinner, M.I. and Rose, E.A., Morphine disrupts nocturnal sleep in a dose-dependent fashion, Anesth. Analg. 68, S200 1989.
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Isono and Rosenberg
185. Cronin, A., Keifer, J.C., Baghdoyan, H.A. and Lydic, R., Opioid inhibition of rapid eye movement sleep by a specific mu receptor agonist, Br. J. Anaesth. 74, 188–192, 1995. 186. Rosenberg, J., Oturai, P., Erichsen, C.J., Pedersen, M.H. and Kehlet, H., Effect of general anesthesia and major versus minor surgery on late postoperative episodic and constant hypoxemia, J. Clin. Anesth. 6, 212–216, 1994. 187. Cronin, A.J., Keifer, J.C., Davies, M.F., King, T.S. and Bixler, E.O., Postoperative sleep disturbance: influences of opioids and pain in humans, Sleep 24, 39–44, 2001. 188. Giubilei, F., Iannilli, M., Vitale, A., Pierallini, A., Sacchetti, M.L., Antonini, G. and Fieschi, C., Sleep patterns in acute ischemic stroke, Acta Neurol. Scand. 86, 567–571, 1992. 189. Broughton, R. and Baron, R., Sleep patterns in the intensive care unit and on the ward after acute myocardial infarction, Electroencephalogr. Clin. Neurophysiol. 45, 348–360, 1978. 190. Hilakivi, I., Biogenic amines in the regulation of wakefulness and sleep, Med. Biol. 65, 97–104, 1987. 191. Fehm, H.L., Benkowitsch, R., Kern, W., Fehm-Wolfsdorf, G., Pauschinger, P. and Born, J., Influences of corticosteroids, dexamethasone and hydrocortisone on sleep in humans, Neuropsychobiology 16, 198–204, 1986. 192. Opp, M., Obal, F., Jr. and Krueger, J.M., Corticotropin-releasing factor attenuates interleukin 1-induced sleep and fever in rabbits, Am. J. Physiol. 257, R528–R535, 1989. 193. Shibasaki, T., Yamauchi, N., Hotta, M., Imaki, T., Oda, T., Ling, N. and Demura, H., Brain corticotropin-releasing hormone increases arousal in stress, Brain Res. 554, 352–354, 1991. 194. Opp, M.R. and Krueger, J.M., Interleukin 1-receptor antagonist blocks interleukin 1-induced sleep and fever, Am. J. Physiol. 260, R453–R457, 1991. 195. Kubota, T., Brown, R.A., Fang, J. and Krueger, J.M., Interleukin-15 and interleukin-2 enhance non-REM sleep in rabbits, Am. J. Physiol. Regul. Integr. Comp. Physiol. 281, R1004–R1012, 2001. 196. Trachsel, L., Schreiber, W., Holsboer, F. and Pollmacher, T., Endotoxin enhances EEG alpha and beta power in human sleep, Sleep 17, 132–139, 1994. 197. Bauer, J., Hohagen, F., Gimmel, E., Bruns, F., Lis, S., Krieger, S., Ambach, W., Guthmann, A., Grunze, H., Fritsch-Montero, R., Weissbach, A., Ganter, U., Frommberger, U., Riemann, D. and Berger, M., Induction of cytokine synthesis and fever suppresses REM sleep and improves mood in patients with major depression, Biol. Psychiatry 38, 611–621, 1995. 198. Kapas, L., Hong, L., Cady, A.B., Opp, M.R., Postlethwaite, A.E., Seyer, J.M. and Krueger, J.M., Somnogenic, pyrogenic, and anorectic activities of tumor necrosis factor-alpha and TNF-alpha fragments, Am. J. Physiol. 263, R708–R715, 1992. 199. Galland, B.C., Bolton, D.P. and Taylor, B.J., Apnea and rapid eye movement sleep excess in the piglet during recovery from hyperthermia, Pediatr. Res. 34, 518–524, 1993.
Recovery from Anesthesia
777
200. Knudsen, J., Duration of hypoxaemia after uncomplicated upper abdominal and thoraco-abdominal operations, Anaesthesia 25, 372–377, 1970. 201. Parfrey, P.S., Harte, P.J., Quinlan, J.P. and Brady, M.P., Postoperative hypoxaemia and oxygen therapy, Br. J. Surg. 64, 390–393, 1977. 202. Singh, N.P., Vargas, F.S., Cukier, A., Terra-Filho, M., Teixeira, L.R. and Light, R.W., Arterial blood gases after coronary artery bypass surgery, Chest 102, 1337–1341, 1992. 203. Bishop, D.G. and McKeown, K.C., Postoperative hypoxaemia: oesophagectomy with gastric replacement, Br. J. Surg. 66, 810–812, 1979. 204. Modig, J., Respiration and circulation after total hip replacement surgery. A comparison between parenteral analgesics and continuous lumbar epidural block, Acta Anaesthesiol. Scand. 20, 225–236, 1976. 205. Hudgel, D.W. and Devadatta, P., Decrease in functional residual capacity during sleep in normal humans, J. Appl. Physiol. 57, 1319–1322, 1984. 206. Series, F. and Marc, I., Influence of lung volume dependence of upper airway resistance during continuous negative airway pressure, J. Appl. Physiol. 77, 840–844, 1994. 207. Leiter, J.C., Knuth, S.L. and Bartlett, D., Jr., The effect of sleep deprivation on activity of the genioglossus muscle, Am. Rev. Respir. Dis. 132, 1242–1245, 1985. 208. Bowes, G., Woolf, G.M., Sullivan, C.E. and Phillipson, E.A., Effect of sleep fragmentation on ventilatory and arousal responses of sleeping dogs to respiratory stimuli, Am. Rev. Respir. Dis. 122, 899–908, 1980. 209. Rosenberg-Adamsen, S., Stausholm, K., Edvardsen, L., Zwarts, M., Kehlet, H. and Rosenberg, J., Body position and late postoperative nocturnal hypoxaemia, Anaesthesia 52, 589–592, 1997. 210. Flemons, W.W., Whitelaw, W.A., Brant, R. and Remmers, J.E., Likelihood ratios for sleep apnea: clinical prediction rule, Am. J. Respir. Crit. Care Med. 150, 1279–1285, 1994. 211. Gold, A.R., Schwartz, A.R., Bleecker, E.R. and Smith, P.L., The effect of chronic nocturnal oxygen administration upon sleep apnea, Am. Rev. Respir. Dis. 134, 925–929, 1986. 212. Martin, R.J., Sanders, M.H., Gray, B.A. and Pennock, B.E., Acute and longterm ventilatory effects of hyperoxia in the adult sleep apnea syndrome, Am. Rev. Respir. Dis. 125, 175–180, 1982. 213. Rosenberg, J., Pedersen, M.H., Gebuhr, P. and Kehlet, H., Effect of oxygen therapy on late postoperative episodic and constant hypoxaemia, Br. J. Anaesth. 68, 18–22, 1992. 214. Reeder, M.K., Goldman, M.D., Loh, L., Muir, A.D., Casey, K.R. and Gitlin, D.A., Postoperative obstructive sleep apnoea. Haemodynamic effects of treatment with nasal CPAP, Anaesthesia 46, 849–853, 1991. 215. Rennotte, M.T., Baele, P., Aubert, G. and Rodenstein, D.O., Nasal continuous positive airway pressure in the perioperative management of patients with obstructive sleep apnea submitted to surgery, Chest 107, 367–374, 1995.
19 Neuromuscular Blocking Agents and Ventilation
LARS I. ERIKSSON Karolinska Hospital and Institute Stockholm, Sweden
In this chapter, the effect of neuromuscular blocking agents (NMBAs) on the regulation of breathing and on the respiratory muscles will be described. The information will be structured as follows:
regulation of breathing; carbon dioxide sensing and NMBAs; oxygen sensing and NMBAs; respiratory pump function and the control of the upper airways; the diaphragm and NMBAs; the upper airways and NMBAs.
Neuromuscular blocking agents (NMBAs) bind reversibly to the ligandgated acetylcholine receptor at the neuromuscular junction and block cholinergic transmission across the synaptic cleft. This cholinergic block is either competitive (nondepolarizing neuromuscular blocking agents) or noncompetitive (depolarizing neuromuscular blocking agents). Although nondepolarizing and depolarizing NMBAs have their preferred affinity at the nicotinic acetylcholine receptor (nAChR) widely distributed in striated skeletal muscles (muscle type nAChR), recent findings demonstrate that 779
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they also bind to the neuronal-type nAChR within the nervous system [1,2]. This results in a number of actions other than those we classically associate with the pharmacological action of neuromuscular blocking agents, i.e., a paralysis of striated skeletal muscles. The initial study providing insight into the effects of curare on respiration was published in 1811–1812 [3]. In a series of experiments, Brodie showed that the lethal effects of curare could be overcome if artificial ventilation were provided to the animal. With great detail after having applied curare to a small wound in the cat, he described the rapid cessation of breathing efforts followed by a gradual return of small respiratory movements, continuing with gross skeletal muscle movements until breathing movements had completely recovered and artificial ventilation was stopped. Unaware of the complexity of the interaction between NMBA and respiratory control, Brodie was the first scientist to demonstrate how NMBAs act on the respiratory muscles. I.
Regulation of Breathing
A. Carbon Dioxide Sensing and NMBAs
Chemosensitivity to carbon dioxide is mainly localized in central respiratory neurons of the brainstem. To a lesser extent, sensitivity to CO2 has been demonstrated within the carotid bodies, thus, carotid body CO2 sensing seems to have a limited impact on normal resting ventilation and regulation of breathing during hypercarbia. Hence, carbon dioxide chemosensation is mainly located within the CNS surrounded by the blood-brain barrier (BBB) (see further Chapters 2 and 15, this volume). NMBAs are generally not thought to interact with central chemoreception. NMBAs are large, bulky molecules built on either a long CHO chain of ester-bindings or an aminosteroidal nucleus. Regardless of the molecular structure upon which the drug is built, the affinity to the muscle type nAChR binding site is due to the action of a positively charged quaternary nitrogen ion at one end of this large molecule. Consequently, the NMBAs are highly hydrophilic compounds with a very poor capacity to pass lipid-containing layers, such as the BBB [4] and the placenta. Consequently, NMBAs are found in negligible amounts in cerebrospinal fluid (CSF), provided the BBB is intact [4,5]. Moreover, placental transfer of NMBAs is negligible and NMBAs do not interfere with breathing movements of the fetus, or breathing in the neonate after administration of NMBAs to the mother. Because of this, it is not surprising that the ventilatory response to hypercarbia (HCVR) is well maintained after administration of subparalysing doses of NMBA [6–8]. In humans, several investigators [6–8] have demonstrated that both the tidal volume and minute volume responses to increased end-tidal CO2 are unchanged during partial neuromuscular block.
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Hence, it is evident that resting ventilation and ventilation during hypercarbia is unaffected by NMBA, provided that the dose is small enough not to affect the diaphragmatic pump function.
B. Oxygen Sensing and NMBAs
In contrast to the central origin of the ventilatory response to carbon dioxide, the acute increase in ventilation during hypoxia is mainly governed by afferent neuronal input from the peripheral chemoreceptors within the carotid bodies and, to some extent, from those of the aortic arch. The peripheral chemoreceptors are thus located within the peripheral nervous system and are not protected by the blood-brain barrier (BBB) from entrance of NMBAs or other hydrophilic compounds. Although the exact nature of carotid body oxygen sensing remains unsolved, it is evident that hypoxia results in a number of cellular events involving multiple ion channels such as calcium and several potassium channels. This ultimately leads to the release of neurotransmitters such as acetylcholine, dopamine, substance P, and ATP; the chemical transmission seem to involve nitric oxide, subsequently leading to depolarization of the carotid sinus nerve. The interplay between these multiple neurotransmitters has not been settled, yet cholinergic transmission seems to play a key role within the carotid body oxygen sensing, both during development (fetal and neonatal life) and in adulthood (for a comprehensive review, see Chapters 1 and 16, this volume, and Ref. [9]). Heymans [10] showed in the 1930s that direct application of acetylcholine onto the carotid bodies resulted in an instant and shortlasting episode of hyperventilation and a simultaneous reduction in heart rate (Figure 19.1). Based on a series of experiments, it was concluded that acetylcholine was involved in transmission of afferent neuronal input from the carotid body to the CNS. For his work on cholinergic transmission of the carotid body [10], Heymans received the Nobel Prize for Medicine in 1939. Although these results have been debated, it is now generally agreed that intact cholinergic transmission is an essential part for normal oxygen sensing [11,12]. Early in vitro studies by Fitzgerald et al. [13] demonstrated that carotid body chemosensitivity, expressed as the increase in whole carotid sinus nerve activity during hypoxia, was completely abolished after administration of D-tubocurarine and atropine. Moreover, nicotinic and muscarinic reactive sites have been shown to be present on the chemoreceptor cell (glomus cell), causing the cell to hyperpolarize after administration of D-tubocurarine [14]. However, acetylcholine does not seem to be the primary neurotransmitter involved in carotid body oxygen sensing; rather, it seems to participate in a modulatory feed-back mechanism. Hence, it seems more likely that multiple transmitters such as
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A 220 200
B
180 160 mm Hg
0.1 mg Acetylcholine
3"
Figure 19.1 (A) Pneumogram of a dog under chloralose anesthesia; (B) systemic blood pressure and heart rate. Time in 3-s intervals. Acetylcholine 0.1 mg was applied to the surface of the carotid bodies, resulting in instant hyperventilation and bradycardia (Data from Ref. 10).
dopamine, acetylcholine, ATP (co-released with ACh), and substance-P are involved in chemical transmission of the carotid body chemoreceptors [15–19] where NO may play a role as second messenger [20]. In 1992, it was reported that a clinically used NMBA (vecuronium) depresses the hypoxic ventilatory response (HVR) in human subjects [21]. The investigation was carried out using a poikilocapnic hypoxic ventilatory test procedure [21] and was later reconfirmed, after similar findings using a normo-isocapnic hypoxic ventilatory test procedure after vecuronium [22] as well as after other clinically used nondepolarizing NMBA [8]. Hence, the acute hypoxic ventilatory response is reduced by approximately 30% after equipotent administration of compounds such as atracurium, vecuronium and pancuronium (Figure 19.2). Moreover, the magnitude of HVR
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HVR mL/min/% SpO2
Hypoxic Ventilatory Response 500
ATRACURIUM
PANCURONIUM
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400 400 300 200 100
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0 Control
TOF 0.70
TOF > 0.90
500
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400
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TOF 0.70
TOF > 0.90
0 Control
TOF 0.70
TOF > 0.90
Figure 19.2 The depression of hypoxic ventilatory response by an NMBA (vecuronium) in volunteers. Hypoxic ventilatory responses (HVR) before (control), during partial neuromuscular block (TOF 0.70), and after recovery (TOF 4 0.90). Data presented as mean S.D.(**)P 5 0.01 (Data from Ref. 8).
depression is the same regardless of molecular structure of the NMBA, whether an aminosteroidal or a bensylisoquinolinium-based compound [8]. Furthermore, the depression is no different in compounds with muscarinic acetylcholine receptor affinity (pancuronium) when compared with those having very low or no muscarinic acetylcholine receptor affinity (vecuronium and atracurium) [8,21,22]. For all drugs, however, it is important to note that there is considerable interindividual variation in the degree of depression of human hypoxic ventilatory responses. Some individuals may have a completely abolished response to hypoxia while others may only have minimal depression [8,22]. It is therefore difficult to predict the depression of HVR in a single individual. It was hypothetized that the underlying mechanism was caused by an interaction between NMBAs and hypoxic chemosensitivity of the carotid bodies [21,22]. In the anesthetized rabbit, it was later shown that injecting small doses of an NMBA (vecuronium) into the carotid body via a lingual artery catheter caused a dose-dependent depression of phrenic nerve activity during systemic hypoxia [23]. Wyon et al. [24] also showed in vivo that the firing frequencies of single isolated chemoreceptors in the carotid body were almost abolished by systemic administration of vecuronium (Figure 19.3). After a complete neuromuscular blocking dose of the NMBA, the depressed chemoreceptor recovered spontaneously. In general, NMBAs have been regarded as having little, if any, affinity to the neuronal-type nAChR as opposed to their target action on the muscle-type nAChR. Since the nAChR population within the carotid bodies is largely of the neuronal type, it can be argued whether the interaction is due to a binding of NMBAs to neuronal-type nAChR or to other receptor types or elements. Recently, Igarashi et al. [25] and Jonsson et al. [26] have shown that the carotid body chemoreceptor responses to nicotine are markedly reduced or even abolished after exposure to NMBAs (Figure 19.4).
CO2, MAP, % mmHg
200 100 5 5
Chemo output, O2, Hz %
25 0
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Eriksson
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Chemo output, O2, Hz %
784
25
0
40 20 0
Control
Vecuronium 0.1 mg
30 min
60 min
Vecuronium 0.5 mg
0 0 40 20 0
90 min
Figure 19.3 Depression of the activity during hypoxia of a single carotid body chemoreceptor after systemic administration of an NMBA. Vecuronium was given i.v. as partial paralysis (0.1 mg), complete paralysis (0.5 mg), and after spontaneous recovery (30, 60 and 90 min). Original recording presented as firing frequencies (Hz) along with arterial mean pressure (MAP) and tracheal O2 and CO2 concentration (%) (Data from Ref. 24).
In addition, Wyon et al. [27] have described the concentration–response relationship for NMBAs and the cholinergic transmission within the carotid bodies, which demonstrates the affinity of NMBAs to the neuronal-type nAChR of the carotid bodies. In summary, neuromuscular blocking compounds interfere with cholinergic transmission of the carotid body, which is most likely the mechanism behind the depression of oxygen sensing. Clearly, this represents a built-in phenomenon (class effect) among NMBAs. The exact nature of this interaction with the carotid body chemoreceptor remains to be clarified. The significance of this interference should be considered. Depressed hypoxic ventilatory responses due to residual effects of NMBAs are a risk factor among postoperative patients. More importantly, the postoperative patient may have further ventilatory depression when an NMBA is used in combination with other potent anesthetic agents (opioids, hypnotics). In addition, in animal experiments, the depression of chemoreceptor activity may have important implications for our interpretation of previous experimental works in the field of carotid body chemoreceptor function.
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Hypoxia Hypoxia
Frequency (Hz)
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Nicotine Nicotine 500µg 500µg
300
Nicotine 500µg
Nicotine 500µg
Nicotine 500µg
200 100
Buffer 25 min
Buffer 25 min
Vecuronium Neo 30 min 30 min
Buffer Buffer 25 min 25 min Recording time (s)
Figure 19.4 The depression of neuronal-type nAChR of the carotid body by an NMBA (vecuronium) in the isolated carotid body preparation [25,26]. Firing frequencies of the whole carotid body sinus nerve after nicotine challenges before and during perfusion with vecuronium and neostigmine. Original recording from one experiment (Data from Ref. 26).
In many studies, NMBAs have been administered as intermittent bolus doses or continuous infusions as part of the anesthetic procedure to keep the animal immobilized. The possibility that this anesthetic technique may have interfered with normal chemoreceptor function, and thus with the results of the experiment, should therefore be kept in mind. Thus, these findings have implications not only for patient care, but also for experimental practice as far as ventilatory studies are concerned.
II.
Respiratory Pump Function and the Control of the Upper Airways
A. The Diaphragm and NMBAs
Respiratory pump function and maintenance of normal gas exchange over a range of workloads are, to a large extent, the work performed by the diaphragm. The contraction pattern of the diaphragm is mainly governed by the efferent activity of central inspiratory and expiratory neurons within the brainstem. The neuromechanical link between these central respiratory neurons and the diaphragm is represented by the two phrenic nerves.
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Neuromuscular blocking agents block neuromuscular transmission at the junction between the phrenic nerve and the diaphragm. Consequently, this results in paralysis of the diaphragm, the degree of paralysis being dependent on the dose of the NMBA administered. Interestingly, the dose needed to produce a 90% depression of diaphragm contraction force (ED90 diaphragm) is about two times higher than that needed for other muscles, i.e., two times the ED90 for peripheral skeletal muscles (hand, leg and abdominal muscles) [28]. Consequently, the diaphragm is resistant to NMBAs when compared with other muscle groups, and normal minute ventilation is well maintained despite considerable weakness of peripheral muscles. In a series of human experiment [6,7], Gal and coworkers described the respiratory sparing effect of NMBAs in volunteers, i.e., they showed that diaphragmatic contraction force (expressed as the capacity to generate negative inspiratory pressures), the resting ventilation and carbon dioxide stimulated ventilation are all well maintained despite almost complete paralysis of peripheral hand muscles (Figure 19.5). Furthermore, during
30
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f 20 (breaths/min)
f
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VT (Litres)
VT 1.0 Control Atropine 0.026 0.043 0.072 0.12 1.0 mg d – Tubocurarine (mg / Kg)
0.20
Figure 19.5 The resistance of the diaphragm to an NMBA (D-tubocurarine). Effects on ventilation during carbon dioxide stimulation. Note the reduction of tidal volumes and the compensatory increase in respiratory frequency, which together maintain minute ventilation unchanged. VE: expired minute ventilation; f: respiratory frequency; VT: tidal volume (Data from Ref. 6).
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more intense neuromuscular block with a subsequent reduction in tidal volume, minute ventilation and end-tidal carbon dioxide tension are kept normal by a compensatory increase in respiratory frequency [7]. Hence, during incomplete neuromuscular block, the resistance of the diaphragm to NMBAs will allow adequate minute ventilation with normal gas exchange, but at the cost of increased work of breathing [6–8]. The time course of action of an NMBA is also different at the diaphragm when compared with other muscle groups. The onset of action of an NMBA is mainly governed by the distribution of the drug, i.e., the blood flow to the target organ. A greater proportion of cardiac output is directed to central respiratory muscle groups than to peripheral muscles. As a result, the distribution time constant (keo) to the effect compartment (i.e., the neuromuscular junction) of the respiratory muscles is considerably greater than that of more peripherally located muscles (e.g., hand, leg). Even with the abovementioned resistance to NMBAs, the onset time to maximum neuromuscular block will therefore be shorter at the diaphragm when compared with peripheral muscle groups [29,30] (see Figure 19.6). Moreover, because of the resistance to NMBAs, the offset of a neuromuscular block is shorter at the diaphragm than at more peripherally located muscle groups [29,30]. 100
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Figure 19.6 The time course of action (evoked twitch force) of three different muscle groups—the diaphragm, the orbicularis oculi and the adductor pollicis (hand)—after a single dose of an NMBA (vecuronium 0.07 mg kg1 i.v.). Note the rapid onset and offset of action in the diaphragm when compared with the adductor pollicis muscle (Data from Ref. 29).
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The muscles that control the upper airways consist of the pharyngeal muscles, the cricopharyngeal segment of the upper esophagus (striated muscle), and the laryngeal elevator and closure muscles. These muscles are far more sensitive to NMBAs than any other peripheral muscle group including the hand, leg, and abdominal muscles [31,32]. In animals, the ED90 is considerably lower for peripheral skeletal muscles and the diaphragm, yet no dose–response data for NMBAs and the upper airway muscles are available for humans. Interestingly, the larynx is, on the other hand, an organ with a sensitivity and time course of action of the NMBAs that is quite similar to that of the diaphragm, making these muscles the exception to this rule [33]. The central location of the upper airway muscles makes them extremely sensitive to even very small doses of an NMBA [31,32, 34–36]. As illustrated by D’Honneur et al. [31], airway obstruction may occur during a partial neuromuscular block [34]. Because of the rapid distribution to this muscle compartment, the onset time of block is short. However, because of their extreme sensitivity to NMBAs, the neuromuscular block will also have a considerably more prolonged time course when compared with that of the diaphragm [30] (see Figure 19.7). Thus, it results in an uneven recovery pattern among the muscles controlling the upper airway when compared with the respiratory pump muscle (diaphragm). Consequently, an incomplete neuromuscular block will have a more pronounced action in the pharyngeal constrictor muscles, the cricopharyngeal muscle and those controlling the laryngeal inlet (Figure 19.8). The differential
Tgh
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Figure 19.7 The time course of action of an NMBA in the diaphragm compared with the pharynx of an anaesthetized dog. Note the more pronounced paralysis in the pharynx (Tgh) as compared with the diaphragm (Pdi) as well the prolonged recovery in the pharyngeal muscle (Data from Ref. 30).
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150
100
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Figure 19.8 The sensitivity of the upper esophageal sphincter muscle compared with the adductor pollicis muscle. Note the marked depression of upper esophageal sphincter resting tension despite almost complete return of force (TOF 0.60–0.80) in the adductor pollicis muscle (Data from Ref. 35).
effects within the pharynx also lead to a dyscoordination between the propelling constrictor muscle group and the upper esophageal segment. The net result leads to an increased risk for aspiration of oral contents into the trachea and failure to maintain the airway [35,36]. This may expose postoperative patients to an increased risk for serious adverse events in the immediate postoperative period from the presence of residual NMBA effects. The reason for the difference in sensitivity between muscle groups remains unsolved. Several explanations and hypotheses have been put forward. Based on recent studies, it seems unlikely that the difference is due to muscle fiber size, muscle fiber type, nAChR density, or to the ratio between fiber size and receptor density [37–39]. It has recently been speculated that a difference in the cholinergic neurotransmitter release pattern between muscles under continued contraction (i.e., sensitive muscles) and those with intermittent work pattern (i.e., resistant muscles) results in a different sensitivity to NMBA [40].
References 1.
Chiodini, P.M., Charpantier, E., Muller, D., Tassonyi, E., Fuchs-Buder, T. and Bertrand, D., Blockade and activation of the human neuronal nicotinic acetylcholine receptors by atracurium and laudanosine, Anesthesiology 94, 643–651, 2001.
790 2.
3.
4. 5.
6.
7.
8. 9.
10.
11.
12.
13.
14. 15.
16. 17.
Eriksson Violet, J.M., Downie, D.L., Nakisa, R.C., Lieb, W.R. and Franks, N.P., Differential sensitivities of mammalian neuronal and muscle nicotinic acetylcholine receptors to general anesthetics, Anesthesiology 86, 866–874, 1997. Brodie, B.C., Experiments and observations on the different modes in which death is produced by certain vegetable poisons, Phil. Trans. Roy. Soc. 101, 194–195, 1811. Matteo, R.S., Pua, E.K., Khambatta, H.J. and Spector, S., Cerebrospinal fluid levels after D-tubocurarine in man, Anesthesiology 46, 396–399, 1977. Werba, A., Gilly, H., Weindlmayr-Goettel, M., Spiss, C.K., Steinbereithner, K., Czech, T. and Agoston, S., Porcine model for studying the passage of non-depolarizing neuromuscular blockers through the blood brain barrier, Br. J. Anaesth. 69, 382–386, 1992. Gal, T.J. and Smith, T.C., Partial paralysis with D-tubocurarine and the ventilatory response to CO2. An example of respiratory sparing? Anesthesiology 45, 22–28, 1976. Gal, T.J. and Goldberg, S.K., Relationship between respiratory muscle strength and vital capacity during partial curarization in awake subjects, Anesthesiology 54, 141–147, 1981. Eriksson, L.I., Reduced hypoxic chemosensitivity in partially paralysed man. A new property of muscle relaxants? Acta Anaesthesiol. Scand. 40, 520–523, 1996. Gonzalez, C., Almaraz, L., Obeso, A. and Rigual, R., Carotid body chemoreceptors: from natural stimuli to sensory discharges, Physiol. Rev. 74, 829–898, 1994. Heymans, C.J.F., The part played by vascular presso- and chemoreceptors in respiratory control, Nobel Lectures – Physiology or Medicine (1922–1941), Amsterdam: Elsevier, pp. 460–481, 1965. Fitzgerald, R.S., Oxygen and carotid body chemotransduction: the cholinergic hypothesis — a brief history and new evaluation, Resp. Physiol. 120, 98–104, 2000. Fitzgerald, R.S., Shirahata, M. and Wang, H.-Y., Acetylcholine is released from in vitro cat carotid bodies during hypoxic stimulation, in Oxygen Sensing: Molecule to Man, Lahiri, S., Prabakhar, N.R. and Forster R.E., eds., New York, Kluwer Academic/Plenum Publishers, pp. 485–493, 2000. Fitzgerald, R.S. and Shirata, M., Carotid body neurotransmission, in Neurobiology and Cell Physiology of Chemoreception, Data, P.G., Acker, H. and Lahiri S., eds., New York, Plenum Press, pp. 131–136, 1991. Eyzaguirre, C. and Monti-Bloch, L., Nicotinic and muscarinic reactive sites in mammalian glomus cells, Brain Res. 252, 181–184, 1982. Kim, D., Oh, E.K., Summers, B.A., Prabakhar, N.R. and Kumar, G.K., Evidence for the involvement of voltage-gated Ca2þ channels in hypoxiainduced release of substance P from carotid body (abstr), FASEB J. 14, A394, 2000. Fitzgerald, R.S., Shirahata, M. and Wang, H.-Y.J., Acetylcholine release from cat carotid bodies, Brain Res. 841, 53–61, 1999. Zhang, M., Zhong, H., Vollmer, C. and Nurse, C.A., Co-release of ATP and ACh mediates hypoxic signaling at rat carotid body chemoreceptors, J. Physiol. 525, 143–158, 2000.
Neuromuscular Blocking Agents and Ventilation 18.
19. 20. 21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
791
Prasad, M., Fearon, I.M., Zhang, M., Laing, M., Vollmer, C. and Nurse, C.A., Expression of P2X2 and PSX3 receptor subunits in rat carotid body afferent neurons in chemsensory signaling, J. Physiol. 537, 667–677, 2001. Prabakhar, N.R., Oxygen sensing of the carotid body chemoreceptors, J. Appl. Physiol. 88, 2287–2295, 2000. Prabakhar, N.R., NO and CO as second messengers in oxygen sensing in the carotid body, Respir. Physiol. 115, 161–168, 1999. Eriksson, L.I., Lennmarken, C., Wyon, N. and Johnson, A., Attenuated ventilatory response to hypoxaemia at vecuronium-induced partial neuromuscular block, Acta Anaesthesiol. Scand. 36, 710–715, 1992. Eriksson, L.I., Sato, M. and Severinghaus, J.W., Effect of vecuronium-induced partial neuromuscular block on hypoxic ventilatory response, Anesthesiology 78, 693–699, 1993. Wyon, N., Eriksson, L.I., Yamamoto, Y. and Lindahl, S.G.E., Vecuroniuminduced depression of phrenic nerve activity during hypoxia in the rabbit, Anesth. Analg. 82, 1252–1256, 1996. Wyon, N., Joensen, H., Yamamoto, Y., Lindahl, S.G.E. and Eriksson, L.I., Carotid body chemoreceptor function is impaired by vecuronium during hypoxia, Anesthesiology 89, 1471–1479, 1999. Igarashi, A., Amagasa, S., Horikawa, H. and Shirahata, M., Vecuronium directly inhibits hypoxic neurotransmission of the rat carotid body, Anesth. Analg. 94, 117–122, 2002. Jonsson, M., Kim, C., Yamamoto, Y., Runold, M.K., Lindahl, S.G.E. and Eriksson, L.I., Atracurium and vecuronium block nicotine induced carotid body responses, Acta Anaesthesiol. Scand. 46, 488–494, 2002. Jonsson, M., Wyon, N., Lindahl, S.G.E., Fredholm, B.B. and Eriksson, L.I., Neuromuscular blocking agents block carotid body neuronal nicotinic acetylcholine receptors, Eur. J. Pharmacol. 497, 173–180, 2004. Lebrault, C., Chauvin, M., Guirimand, F. and Duvaldestin, P., Relative potency of vecuronium on the diaphragm and the adductor pollicis, Br. J. Anaesth. 63, 389–392, 1989. Donati, F., Meistelman, C. and Plaud, B., Vecuronium neuromuscular blockade at the diaphragm, the orbicularis oculi, and the adductor pollicis muscles, Anesthesiology 73, 870–875, 1990. Isono, S., Koichi, T., Ide, T., Suigimori, K., Mizuguchi, T. and Nishino, T., Differential effects of vecuronium at the diaphragm and geniohyoid muscle in anesthesized dogs, Br. J. Anaesth. 68, 339–342, 1992. D’Honneur, G., Gall, O., Gerard, A., Pimaniol, J.M. and Lambert, Y., Priming doses of atracurium and vecuronium depress swallowing in humans, Anesthesiology 77, 1070–1073, 1992. Pavlin, E.G., Holle, R.H. and Schoene, R., Recovery of airway protection compared with ventilation in humans after paralysis with curare, Anesthesiology 70, 381–385, 1989. Wright, P.M., Caldwell, J.E. and Miller, R.D., Onset and duration of rocuronium and succinylcholine at the adductor pollicis and laryngeal adductor muscles in anesthetized humans, Anesthesiology 81, 1110–1115, 1994.
792 34.
35.
36.
37.
38.
39.
40.
Eriksson D’Honneur, G., Lofaso, F., Drummond, G.B., Rimaniol, J.-M., Aubineau, J.V., Harf, A. and Duvaldestin, P., Susceptibility to upper airway obstruction during partial neuromuscular block, Anesthesiology 88, 371–378, 1998. Eriksson, L.I., Sundman, E., Olsson, R., Nilsson, L., Witt, H., Ekberg, O. and Kuylenstierna, R., Functional assessment of the pharynx at rest and during swallowing in partially paralyzed humans, Anesthesiology 87, 1035–1043, 1997. Sundman, E., Witt, H., Olsson, R., Ekberg, O., Kuylenstierna, R. and Eriksson, L.I., The incidence and mechanisms of pharyngeal and upper esophageal dysfunction in partially paralyzed humans, Anesthesiology 92, 977–984, 2000. Ibenbunjo, C., Srikant, C.B. and Donati, F., Morphological correlates of the differential responses of muscles to vecuronium, Br. J. Anaesth. 83, 284–291, 1999. Ibenbunjo, C., Srikant, C.B. and Donati, F., Duration of succinylcholine and vecuronium blockade but not potency correlates with the ratio of endplate to fiber size in seven muscles in the goat, Can. J. Anaesth. 43, 485–494, 1996. Sundman, E., Yost, C.S., Margolin, G., Kuylenstierna, R., Ekberg, O. and Eriksson, L.I., Acetylcholine receptor density in human cricopharyngeal muscle and pharyngeal constrictor muscle, Acta Anaesthesiol. Scand. 46, 999–1002, 2002. Sundman, E., Pharyngeal Function and Anesthetic Agents, Medical Dissertation, Karolinska Institute, Stockholm, Sweden, 2001.
20 Cardiovascular Drugs and the Control of Breathing
DENHAM S. WARD and SUZANNE KARAN University of Rochester School of Medicine and Dentistry Rochester, New York
I.
Introduction
Since coordination of the cardiovascular and respiratory systems is needed in order to optimize oxygen transport, it is not surprising that drugs used to treat cardiovascular disease may have respiratory effects. Because of the close coupling between these systems, it is important to separate the respiratory effects of direct (or reflex) drug actions from those secondary to the improvement in the cardiovascular disease being treated. Thus, the respiratory effects of a drug when tested in normal subjects may be different from the clinical effects seen when the same drug is used in patients. The interactions between the cardiac and respiratory systems were studied by Leonardo da Vinci [1], but a knowledge of respiratory gases and circulation of blood was needed before there could be any real understanding of the close coupling between the two systems. The study of pharmacological interaction in the cardio-respiratory system may have started in the late 19th century, when researchers in London observed that exogenously administered suprarenal extract reduced minute ventilation [2]. Some years later, another research team found that 793
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lowering blood pressure by inducing hemorrhage in cats caused an increase in minute ventilation, establishing a direct physiological link between the respiratory and cardiovascular systems [3]. The relationships between peripheral chemoreceptors and the cardiovascular system have recently been reviewed [4]. While many of the respiratory effects of cardiovascular drugs are of physiologic interest, few such drugs are limited clinically by their respiratory side effects. In fact, the benefits of most cardiovascular drugs in maintaining homeostasis usually far outweigh any direct negative respiratory effects. Many of these respiratory effects are only seen under laboratory conditions using animal models or normal subjects, and therefore many of the respiratory effects of cardiovascular drugs are interesting because of the insight they provide into the pharmacology of the control of breathing. Regulation of the cardiovascular and respiratory systems shows a strong coordination, particularly in response to a stimulus such as hypoxia, which activates both systems. Much of this coordination takes place via brainstem centers that are active in both cardiovascular and respiratory regulation. As stated above, changes in blood pressure directly lead to changes in respiration [5], and there is a central interaction between the baroreceptor and chemoreceptor reflexes [6,7]. Respiratory sinus arrhythmia is another sign of cardio-respiratory coupling that is useful for assessing vagal cardiac input [8]. It has long been appreciated that changes in cardiac output can greatly affect ventilatory stability [9], and it is interesting that treatment of nocturnal bradycardia via pacing can ameliorate the symptoms of obstructive sleep apnea [10]. A review of the interconnected physiology of cardiac and respiratory control is beyond the scope of this chapter; a volume on this topic has been recently published in this series [11].
II.
Catecholamine Agonists and Antagonists
Since catecholamines are important neurotransmitters and neuromodulators in the autonomic nervous system, catecholamine agonists and antagonists are important therapeutic agents in cardiovascular disease and can have important effects on ventilatory control. One of the earliest respiratory pharmacology studies examined the effects of adrenalin on respiration [12], and the effects of catecholamines on breathing have been reviewed more recently [13–15]. Table 20.1 summarizes the cardiovascular effects of common currently used clinical catecholamine agonists. The catecholamines (both endogenous and exogenous) most studied with respect to their effects on ventilation
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Table 20.1 Comparison of physiological effects of clinically used catecholamines (Data from Ref. 46). Dopamine
Cardiac output Systemic blood pressure Heart rate Systemic vascular resistance Renal blood flow Glomerular filtration rate Chemoreceptor function A-aPo2 O2 transport
Norepinephrine
Low-Dose (55 mg kg1 min1)
High-Dose (45 mg kg1 min1)
Dobutamine
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A-aPo2 indicates alveolar-arterial O2 tension difference; $, no change; ", increase; #, decrease; "", more profound increase; ##, more profound decrease.
are dopamine, epinephrine and norepinephrine. The drugs that affect ventilation via an adrenergic mechanism act primarily at D2, a1 and a2, and b1 and b2 receptor subtypes; drugs with receptor and subtype specificity can provide additional information about respiratory and cardiovascular interactions. The prototypical adrenergic drugs norephinephrine (NE) and epinephrine (E) are still used clinically and act relatively nonspecifically on both a and b receptors. They differ primarily in that NE has less activity at b2 receptors [16]. Therefore, the pharmacological effect of these drugs is a combination of their direct receptor activity together with the activation of vascular and other reflexes. The separation of direct and indirect effects is difficult since, for example, epinephrine can increase both carbon dioxide production and plasma Kþ, both of which are ventilatory stimulants. Linton et al. concluded that all of the carotid body stimulating effects of epinephrine could be accounted for by these mechanisms [17]. In the newborn piglet, epinephrine reversed the ventilation depression caused by hypoxia (hypoxic ventilatory decline, HVD) and also prevented the decrease in oxygen consumption seen with 10 min of hypoxia [18]. Both of these effects required intact carotid bodies.
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The episodic hypoxemia of sleep apnea can elevate plasma NE levels [19], and NE has also been shown to have a direct effect on ventilation. Intracarotid administration of NE depressed ventilation in goats [20] and intravenous NE caused similar results in dogs [21]. In fact, in the latter study, even mechanically ventilated dogs had a NE-induced fall in arterial PO2 and rise in arterial PCO2. A. Dopaminergic Agonists and Antagonists
Dopamine was extensively studied initially in the field of neurobiology, where it was first thought to be primarily an intermediate in the synthesis of norepinephrine and epinephrine; however, it was soon appreciated as a separate neurotransmitter [22] with cardiovascular effects [23,24]. It was also identified as the primary catecholamine in the carotid body type-1 cells of many species (see Nurse, Chapter 1, this volume and the recent review by Gonzalez et al. [25]). Clinically, the drug has traditionally been used and understood in the context of a cardiovascular and renal drug, but it also has clinically important respiratory properties. When given at low doses (55 mg kg1 min1), intravenous dopamine preferentially activates dopaminergic receptors (both D1 and D2), while at high doses it additionally activates both a- and b-adrenergic receptors, causing an increase in systemic vascular resistance and cardiac contractility. Usual intravenous doses do not cross the blood–brain barrier in significant amounts, so only the peripheral effects are apparent even though dopamine is also an important central neurotransmitter in ventilatory control [26]. Though low-dose dopamine has been traditionally advocated for clinical use because it causes increased renal blood flow, activation of peripheral dopaminergic receptors at the carotid bodies may be a significant side effect [27], since exogenous dopamine appears to be a ventilatory depressant in mammals and in humans [28–32]. It has long been appreciated that dopamine inhibits the carotid sinus nerve, perhaps accounting for the hypoventilation seen during its administration [31,33]. However, many studies have also demonstrated the ventilatory depressant effects of dopamine in the absence of carotid bodies in several species [28–30]. There is incomplete elucidation of the mechanism whereby dopamine interacts with ventilation. In fact, it may not always be a direct respiratory effect, but rather an indirect effect resulting from its action on vascular smooth muscle. Loos et al. [34] studied dopamine inhibition of ventilation in cats, differentiating the ventilatory effects of low versus high doses of the drug. Low-dose dopamine suppressed activity of the carotid bodies, while at high doses, the increased blood pressure also caused a baroreceptormediated ventilatory depression. There also seemed to be an a1 receptor vasoconstriction-mediated depression, perhaps through stimulation of group III and IV muscle receptors [34]. Studies have demonstrated that a
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Figure 20.1 The effect of low-dose dopamine (5 mg kg1 min1) on ventilation, saturation and end-tidal CO2 in patients with congestive heart failure (Data from Ref. 36).
low-dose dopamine infusion, given during hypoxia or during states with compromised oxygen delivery to tissues (e.g., congestive heart failure), has depressant effects on minute ventilation (Figure 20.1) [35,36]. Low-dose dopamine has been shown to impair the ventilatory response to hypoxemia and hypercapnia [32], to reduce the isocapnic hypoxic sensitivity by 50–80% [37], and to reduce hypercapnic sensitivity in cats [38] and in humans [39]. This influence is mediated primarily through exogenous dopamine affecting dopamine receptors in the carotid body, and not via a- or b-receptor modulation [32]. Dopamine also attenuates the positive interaction between hypoxia and hypercapnia at the carotid body [40]. Furthermore, it acts centrally to attenuate the declining phase of HVD [41], but ordinarily intravenous dopamine does not cross the blood-brain barrier in sufficient concentration to have a central effect. Laboratory investigations of dopaminergic influences on ventilation have furthered our understanding of the control of ventilation. The quandary of the role of endogenous dopamine’s inhibitory effects on the carotid body, especially in the face of hypoxia, has remained unresolved [25]. It is possible that adjustments in the gain of the fast-responding feedback sensor are required to prevent the oscillations seen in the ventilatory control system (i.e., Cheyne-Stokes breathing [42,43]). This is supported by Lahiri et al. who found that increasing the peripheral chemoreceptor gain with dopamine blockade via domperidone augmented these ventilatory oscillations [44]. Dopaminergic mechanisms may also underlie the decrease in chemosensitivity sometimes seen in patients with obstructive sleep apnea [45]. It is important to watch for signs of sleep apnea when ICU patients are weaned from mechanical ventilation while still receiving a dopamine infusion [46].
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There are other mechanisms whereby dopamine affects respiration. Studies have indicated there is dopaminergic modulation of upper airway muscle tone [47] in a manner that might promote obstruction via the internal pharyngeal constrictors. Inhaled aerosolized dopamine has induced bronchodilation in humans having an asthma crisis, though it does not seem to modulate resting bronchial tone [48], and D2 receptors may be involved in pulmonary C-fiber sensitivity (see Nishino, Chapter 7, this volume) [49]. Fenoldopam is a selective D1-dopamine agonist with moderate a2-receptor affinity, and has been recognized particularly as a vasodilator in the treatment of hypertensive emergencies and for its potential to preserve renal function [50]. Although there are central D1 receptors, intravenous fenoldopam does not cross the blood-brain barrier. Little has been reported about the drug’s direct effects on ventilation, but in rats it has increased lung water reabsorption in a manner similar to dopamine and isoproterenol, presumably through activation of peripheral D1-receptors [51]. Since the inhibitory receptors in the carotid body are D2, fenoldopam would not be expected to depress the hypoxic ventilatory response, but this has not been studied directly. Dopamine antagonists have no useful important cardiovascular effects (and may even be arrhythmogenic [52]) but, as expected, they reverse the dopaminergic inhibition of the carotid bodies [53]. Most dopamine antagonists cross the blood-brain barrier and are used clinically for their sedative, anti-psychotic and anti-emetic properties (see Gross, Chapter 14, this volume). Domperidone, a specific dopamine D2 antagonist that does not cross the blood-brain barrier, when given during hypoxia, increases minute ventilation. The site of action appears to be at D2 receptors on the carotid body [54]. B. a-adrenergic Agonists and Antagonists
Unselective a-adrenergic agonists (e.g, epinephrine, norepinephrine, and phenylephrine) have been traditionally employed as vasoconstrictive agents for their potent peripheral effects. However, the development of selective central a2 -adrenergic agonists has expanded their role for anesthetic [55], sedative [56], and anti-hypertensive effects [57]. Since a2-adrenergic receptor sites are present throughout the central nervous system, and specifically in the brainstem [58], it is a reasonable assertion that they would interact with the control of breathing. O’Halloran et al. studied adrenergic stimulation in awake goats and found that a2 blockade caused modulation of respiratory rhythm during isocapnic hypoxia and potentiated the hypoxic ventilatory response [59]. Furthermore, as the time course and degree of ventilatory potentiation was not changed in carotid body-denervated animals, it appeared that the central component of the response was not modified by peripheral chemoreceptor input during normoxia. This correlates with many
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studies that found the central noradrenergic system to be inhibitory on ventilation through a2 agonism [56,60–62]. However, there have been varying results from studies that examined selective a2 blockade as well as differing theories about the mechanism of central a2-blockade-mediated ventilatory excitation; the contribution of peripheral chemoreceptors cannot be discounted [59]. For example, the carotid body contains a2 receptors [63] and their stimulation has been shown to depress ventilation [64]. It is not clear whether the phenomenon of post-hypoxic frequency decline is dependent on the integrity of central a2 receptors [65]. Recent work has demonstrated that the anti-hypertensive agent clonidine (an a2 agonist) augments hypoxic chemosensory activity through its agonist effects on I1-imidazoline carotid body receptors, in contrast to the inhibitory action of a2-adrenergic agonism at the carotid body [66]. Dexmedetomidine, another a2 agonist used clinically for its analgesic and sedative but not its cardiovascular effects, decreases ventilation in a dose-dependent manner in rabbits and dogs [67,68]. In humans, though, the ventilatory depression and the hypercapnic response associated with intravenous dexmedetomidine administration differed little from placebo control [56]. Thus, in humans, the central a2-agonist effects may change ventilation in relation to their sedative effects and may not directly alter awake ventilation [69]. The ventilatory abnormalities that arise with druginduced sedation can take the form of central apnea or upper airway obstruction. Clonidine given intravenously, orally, and epidurally has been shown to cause apnea [60,61]. Although in one human study even high-dose dexmedetomidine induced no clinically significant apneas [68], it would seem prudent to use this drug cautiously, especially in those patients with risk factors for obstructive sleep apnea. Although a2 receptors exist in the carotid body, the effect of dexmedetomidine on the human hypoxic ventilatory response has not been elucidated. In dogs, even high-dose dexmedetomidine did not alter the hypoxic response [67]. Hedrick et al., studying a2-adrenergic-induced respiratory arrhythmias in awake goats, wondered whether these adrenergic effects were mediated through expiratory-related motoneurons [70]. In later studies, they observed that clonidine did not cause a reduction in the central CO2 drive. Instead, they found that the apneas and respiratory arrhythmias resulted from laryngeal motoneuron activation. An incidental finding was that central CO2 and carotid body chemoreceptors acted to modulate phrenic and recurrent laryngeal nerve activities [70]. C. b-adrenergic Agonists and Antagonists
Central b-adrenergic stimulation facilitates ventilation, probably through b1 actions [71]. Isoproterenol, a nonspecific b-adrenergic agonist, increased ventilation via a carotid body mechanism in the cat [72]. Conversely,
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beta-blockade may inhibit the human ventilatory response to increased sympathetic activity via increased circulating NE or direct NE-stimulation of the CB [73,74]. Further, hypertensive patients have been shown to have a diminished b-adrenergic response in the face of hypoxia [70,75]. There seems to be no evidence that b-receptor-mediated activity modulated the changes occurring during prolonged isocapnic hypoxia [76,77]. III.
Renin–Angiotensin System
The renin-angiotensin system is stimulated during increased ventilatory drive, as occurs during exercise [78] and acute hypercapnia [79]. Heistad et al. found that hypertension caused by an angiotensin infusion was also associated with a decrease in ventilation [73]. Moreover, it seems that both the renal and brain renin-angiotensin systems are involved in modulating respiratory control [80], and there is also evidence for a local angiotensin system in the rat carotid body [81]. Rat carotid bodies have a high density of angiotensin receptors (AT-1) [82] and also contain angiotensinogen mRNA (but not mRNA for renin) [81], but their in vitro response to angiotensin II (AG II) is quite mild [82]. However, with chronic hypoxia, there was an increased density of AT-1 receptors and the in vitro response to AG II was greatly augmented [83]. In the last 15 years, much research has confirmed the role of circumventricular organs (CVO) in regulating respiratory control. These CVO have direct connections with vasopressin V1-receptor sites located in the paraventricular nucleus (PVN) and indirect connections with medullary respiratory centers via neuronal projections from the PVN [79,84]. The role of the CVO in respiration has been confirmed in anesthetized rats [85]. Experimentally, it has been difficult to separate the individual respiratory effects of arginine vasopressin (AVP) and angiotensin II. AVP inhibits the AG II drive to breathe in conscious, normoxic dogs during both eucapnia [86] and hypercapnia [87]. However, prolonged infusions (44 h) of an AG II-antagonist in resting, conscious dogs did not affect respiratory control [88]. In conscious rats, bolus doses of AVP caused depression of minute ventilation and tidal volume, and boluses of AG II caused a reduction in tidal volume, all in the immediate post-infusion period [89]. However, these effects were short-lived, despite the persistent elevation of mean arterial pressure, and the ventilatory parameters returned to baseline by 10 min. After infusing nitroprusside, there was also an immediate increase in respiratory frequency and minute ventilation, which was also short-lived, returning to baseline within 10 minutes. This supports the baroreflex-mediated fast-adapting changes in respiration as a consequence of these drugs, and not their direct ventilatory action [89].
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Baroreceptor-mediated changes in ventilation from AVP infusion returned to baseline after normalization of blood pressure [90]. Interestingly, though, AVP had the independent effect of depressing the slope of the ventilatory response to PCO2 increases [90]. However, there was not the same change in ventilation corresponding to an equivalent AG II-induced rise in mean arterial pressure [90]. Instead, the respiratory effects of AG II seem to be centrally mediated, probably by a CVO mechanism [85]. There also seems to be a role for AG II in modulating the adaptation to prolonged hypoxia in conscious dogs [91], although it does not interact with hypercapnia to affect respiratory control [79]. The role of AVP and AG II in ventilation is further complicated by the current lack of knowledge about the roles plasma osmolality and hormonal imbalance play in the up- and down-regulation of the receptors within the renin-angiotensin system. The genotype of the animal may also play an important role as AG II-receptor blockade depressed ventilation in spontaneously hypertensive (SH) rats but not in the control Wistar-Kyoto rats [92]. However, the SH rat had a higher resting metabolic rate and ventilation. Even though modification of the angiotensin system through angiotensin converting enzyme (ACE) inhibition and direct antagonism at angiotensin II receptors has considerable clinical utility in hypertension and congestive heart failure, there is little work relating animal experiments to humans. It is interesting to note that a polymorphism of the ACE gene has been identified in which an insertion (I) allele (rather than the deletion allele) is associated with lower serum and tissue ACE activity [93,94]. This I allele seems to be associated with improved exercise performance and with muscle response to training [93,94]. Elite mountaineers with the II genotype maintained a higher saturation with rapid (but not slow) ascent to altitude [95]. Differences in sea level acute isocapnic hypoxic ventilatory responses were not reported in this study. Thus, further work is needed to define the role in ventilatory control of the renin-angiotensin system and its pharmacological alteration. IV.
Calcium Channel Blockers
The influx of Ca2þ into the cytosol is a vital step in muscle excitationcontraction coupling and in excitation-secretion coupling in neurons and exocytotic cells [96]. In the carotid bodies, the release of neurotransmitters from type 1 cells in response to hypoxia and hypercapnia involves the opening of membrane calcium channels (see Nurse, Chapter 1, this volume; Lopez-Barneo et al. [97] and Gonzalez et al. [25]). Calcium channel blockers are frequently used clinically in the treatment of hypertension, angina and certain cardiac arrhythmias as well as in the treatment of high-altitude pulmonary edema [98].
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L, N and P/Q types of voltage-gated Ca2þ channels have been identified using specific toxins and, more recently, with studies of the molecular structure of the receptor subunits. Studies of carotid body type-1 cells have identified L-, N- and P-type channels as well as other Ca2þ currents that are resistant to traditional blockers [99]. L-type Ca2þ blockers have been found to inhibit at least part of the hypoxia-sensitive Ca2þ current in rats [90] and rabbits [90] as well as to inhibit neurotransmitter release in rabbits [100]. In dial-urethane anesthetized rats, verapamil caused a dose-related reduction in the hypoxic ventilatory response [101], but in human studies, verapamil appears to have little effect on either the acute or the sustained hypoxic ventilatory response [102,103]. However, following 25 min of hypoxia, verapamil attenuated the increase in ventilation following the transient withdrawal of the hypoxic stimulus [103]. The differences between cellular, animal and human studies could reflect anesthesia effects, species differences, and channel-blocker specificity. An interesting example of complex respiratory-related drug interactions is the study of the effects of Ca2þ blockers on the analgesic effects of morphine. It has been noted in both animals and humans that Ca2þ blockers apparently potentiate the analgesic effects of morphine [104]. In rats, there is an indication that although respiratory depression is increased by the combination of nimodipine and sufentanil, the depression was increased to a lesser extent than the increase in analgesia [105]. In rhesus monkeys, diltiazem enhanced the analgesic potency of morphine but not that of fentanyl [106], and diltiazem did not increase the respiratory depression caused by morphine. Whether this interaction is pharmacodynamic or pharmacokinetic is uncertain (see Olofsen and Dahan, Chapter 5, this volume, for a more complete discussion of the problems inherent in studying drug interactions). V.
Purinoceptor Agonists and Antagonists
The purine nucleoside adenosine is present in both central and peripheral neural tissue where it seems to have properties of a neuromodulator rather than of a classic neurotransmitter. Adenosine, because of its actions on the cardiac conducting system (SA and AV nodes), is used clinically as a fast intravenous bolus to terminate rapid supraventricular tachycardia. While the drug’s respiratory effects are not of major clinical significance in this acute setting (pharmacological cardioversion), its respiratory effects are nonetheless significant. Interestingly, like dopamine, its central effects are the opposite of its peripheral effects, but here, adenosine seems to stimulate peripheral chemoreceptors while depressing central respiratory centers. Clinically used drugs related to adenosine include dipyridamole, an intracellular uptake blocker, used for its chronic platelet inhibitory and
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vasodilating properties; and theophylline (and its water soluble analog, aminophylline), an adenosine-receptor antagonist, used for its bronchodilating properties in asthmatics and for its ability to increase diaphragmatic contractility in ICU patients [107]. When adenosine is given as a bolus in humans, it causes a variety of subjective symptoms (e.g., chest and neck tightness, facial flushing), an increase in breathing frequency and tidal volume, and a transient bradycardia [108]. These effects last only a few seconds because of the rapid intravascular metabolism of adenosine. In a more complete study using an infusion up to 80 mg kg1 min1, which caused no subjective symptoms, Maxwell et al. found a 21% increase in ventilation with concurrent hypocapnia and a small increase in heart rate [109]. They also found an increase in the hypoxic ventilation response (from 0.68 0.4 l min1 %sat1 control to 2.4 1.2 with adenosine) but no change in the hypercapnic response, measured with a progressive isocapnic hypoxic and hyperoxic hypercapnic stimulus, respectively. Since adenosine does not readily cross the blood-brain barrier, these effects most likely arise from peripheral chemoreceptors. A
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Figure 20.2 Effect of the adenosine antagonist, dipyridamole, on the ventilatory response to hypoxia in a single subject. Left panel shows the control response to an isocapnic hypoxic step and the right panel is following dipyridamol, 0.5 mg kg1 i.v., pre-treatment. Both the acute response and the subsequent decline were increased (Data from Ref. 110).
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However, dipyridamole, which inhibits the cellular uptake of adenosine and thus increases its extracellular concentration, does cross the blood-brain barrier. When dipyridamole was given to human subjects, Yamamoto et al. found an increase in the initial acute hypoxic response and also an increase in the ventilatory decline with sustained hypoxia (Figure 20.2) [110]. This effect was attenuated by pretreatment with aminophylline, an adenosine antagonist. These results indicate a peripheral stimulatory and central depressing role for adenosine. In animal studies, adenosine is a potential candidate for a central neuromodulator that mediates hypoxic ventilatory decline, since brain adenosine levels increase with hypoxia and since adenosine has a depressant effect on central neurons. In fact, pretreatment with aminophylline does reduce the magnitude of HVD [111,112]. However, in humans, HVD occurs primarily with a decrease in tidal volume and with no change in respiratory rate [113]. Following aminophylline, ventilation remains constant with sustained hypoxia but with a progressive increase in respiratory rate offsetting the decline in tidal volume that is still present. Thus, central adenosine may not be the direct cause of HVD. In rats, using amperometric enzymatic sensors, Gourine et al. found that the time course of adenosine increase in the nucleus tractus solitarii was too delayed to cause the observed HVD [114]. Periodic breathing with central apneic episodes (Cheyne-Stokes respiration) has long been noted in association with congestive heart failure and systolic dysfunction [9]. Javaheri et al. noted that oral theophylline reduced the number of central (but not obstructive) apneic episodes and also reduced the percent of the night spent in hypoxia in patients with wellcompensated heart failure [115]. They speculated this was due to adenosine antagonism and stimulation of respiration without any increase in hypercapnic sensitivity [107]. VI.
Other Agents
As mentioned in the introduction, hypotension can directly affect chemoreceptors, although it is difficult to separate a direct effect from those reflex effects mediated by the baroreceptors and a central interaction [6]. As a control for the blood pressure effect of other neuromodulators, Maxwell et al. [116] lowered the blood pressure in human subjects with nitroprusside, finding an increase in resting ventilation and an augmentation of hypoxic response without any change in hypercapnic response. Although this effect was ascribed to alterations of blood pressure and local blood flow within the carotid body, it is now known that NO is a neuromodulator or second messenger in the carotid body [117,118]. Alteration in carotid body NO was possibly induced by a
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nitroprusside infusion [119,120], but the increase in local NO caused by nitroprusside might be expected to inhibit the hypoxic response [121]. However, NO may play a dual role in the carotid body, with both stimulation and inhibition depending on its effect, either on the Type 1 cell or on the blood vessels [122]. Nitroprusside also reduces the inhibition of cat carotid body sensitivity to dopamine, but this effect has not been confirmed in humans [123]. There may also be central effects of increased NO on the hypoxic response [124]. Digitalis is probably the oldest therapeutic agent used in cardiac disease. Although it has a very low therapeutic ratio, it is still a useful drug for controlling the ventricular rate in atrial fibrillation, particularly in patients with heart failure. Because it is such an old drug, it has escaped some of the intense scrutiny given to newer agents. McQueen and Ribeiro extensively studied a related drug, oubain, in cats and found that intracarotid injection initially increased chemoreceptor discharge (after several minutes’ delay) but with prolonged infusion, the carotid bodies were desensitized to hypoxia [125]. However, digitalis, given orally as a single dose to normal subjects, seems to augment the acute hypoxic response without altering baseline or hypercapnic ventilation [126]. This could be related to the blockade of sodium-potassium ATPase in the carotid body by digoxin, thereby increasing the excitability of type 1 cells. A single dose may not be sufficient to cause the desensitization seen by McQueen and Ribeiro [125]. References 1.
2. 3. 4. 5. 6.
7.
Mitzner, W.A., Leonardo and the physiology of respiration, in A History of Breathing Physiology, Proctor, D.F., ed., New York, Marcel Dekker, pp. 37–59, 1995. Oliver, G. and Schafer, E.A., The physiological effects of extracts of the suprarenal capsules, J. Physiol. (Lond.) 18, 230–276, 1895. D’Silva, J.L., Gill, D. and Mendel, D., The effects of acute haemorrhage on respiration in the cat, J. Physiol. (Lond.). 187, 369–377, 1966. Marshall, J.M., Peripheral chemoreceptors and cardiovascular regulation, Physiol. Rev. 74, 543–594, 1994. Freyschuss, U. and Melcher, A., Respiratory sinus arrhythmia in man: relation to cardiovascular pressures, Scand. J. Clin. Lab. Invest. 36, 221–229, 1976. Heistad, D.D., Abboud, F.M., Mark, A.L. and Schmid, P.G., Interaction of baroreceptor and chemoreceptor reflexes. Modulation of the chemoreceptor reflex by changes in baroreceptor activity, J. Clin. Invest. 53, 1226– 1236, 1974. Heistad, D., Abboud, F.M., Mark, A.L. and Schmid, P.G., Effect of baroreceptor activity on ventilatory response to chemoreceptor stimulation, J. Appl. Physiol. 39, 411–416, 1975.
806 8.
9. 10.
11. 12. 13.
14.
15.
16.
17.
18.
19. 20.
21.
22.
Ward and Karan Taha, B.H., Simon, P.M., Dempsey, J.A., Skatrud, J.B. and Iber, C., Respiratory sinus arrhythmia in humans: an obligatory role for vagal feedback from the lungs, J. Appl. Physiol. 78, 638–645, 1995. Javaheri, S., A mechanism of central sleep apnea in patients with heart failure, N. Engl. J. Med. 341, 949–954, 1999. Garrigue, S., Bordier, P., Jais, P., Shah, D.C., Hocini, M., Raherison, C., Tunon De Lara, M., Haissaguerre, M. and Clementy, J., Benefit of atrial pacing in sleep apnea syndrome, N. Engl. J. Med. 346, 404–412, 2002. Scharf, S.M., Pinsky, M.R. and Magder, S., Respiratory–Circulatory Interactions in Health and Disease, New York, Marcel Dekker, 2001. Nice, L.E., Rock, L.J. and Courtright, R.O., The influence of adrenalin on respiration, Am. J. Physiol. 34, 326–331, 1914. Mueller, R.A., Lundberg, D.B.A., Breese, G.R., Hedner, J., Hedner, T. and Jonason, J., The neuropharmacology of respiratory control, Pharmacol. Rev. 34, 255–285, 1982. Dempsey, J.A., Olson, E.B. Jr. and Skatrud, J.B., Hormones and neurochemicals in the regulation of breathing, in Handbook of Physiology. The Respiratory System, Vol. II, Fishman, A.P., Cherniack, N.S., Widdicombe, J.G. and Geiger S.R., eds., Bethesda, American Physiological Society, pp. 181– 221, 1986. Ward, D.S. and Temp, J.A., Neuropharmacology of the control of ventilation, in Anesthesia: Biologic Foundations, Biebuyck, J., Lynch, C., Maze, M., Saidman, L.J., Yaksh, T.L. and Zapol, W., eds., Philadelphia, LippincottRaven, pp. 1367–1394, 1997. Hoffman, B.B., Catecholamines, sympathomimetic drugs, and adrenergic receptor antagonists, in The Pharmacological Basis of Therapeutics, Hardman, J.G., Limbird, L.E., eds., New York, McGraw-Hill, pp. 215–268, 2001. Linton, R.A., Band, D.M. and Wolff, C.B., Carotid chemoreceptor discharge during epinephrine infusion in anesthetized cats, J. Appl. Physiol. 73, 2420– 2424, 1992. Suguihara, C., Hehre, D. and Bancalari, E., Effects of epinephrine on the cardiorespiratory response to hypoxia in sedated newborn piglets with intact and denervated carotid bodies, Biol. Neonate 67, 352–359, 1995. Eisenberg, E., Zimlichman, R. and Lavie, P., Plasma norepinephrine levels in patients with sleep apnea syndrome, N. Engl. J. Med. 322, 932–933, 1990. Pizarro, J., Warner, M.M., Ryan, M., Mitchell, G.S. and Bisgard, G.E., Intracarotid norepinephrine infusions inhibit ventilation in goats, Respir. Physiol. 90, 299–310, 1992. Millis, R.M., Trouth, C.O., Johnson, S.M., Wood, D.H., Dehkordi, O. and Wray, S.R., Arterial oxygen desaturation and respiratory failure associated with sympathetic overactivity, in Ventral Brainstem Mechanisms and Control of Respiration and Blood Pressure, Trouth, C.O., Millis, R.M., Kiwull-Schone, H. and Schlafke, M., eds., New York, Marcel Dekker, 1995. Dopamine, in The Biochemical Basis of Neuropharmacology, Cooper, J.R., Bloom, F.E. and Roth, R.H., eds., Oxford, Oxford University Press, pp. 225–270, 2003.
Cardiovascular Drugs and the Control of Breathing 23.
24. 25. 26.
27. 28. 29. 30.
31.
32. 33.
34. 35.
36. 37. 38.
39. 40. 41. 42.
807
Goldberg, L.I. and Sjoerdsma, A., Effects of several monoamine-oxidase inhibitors on the cardiovascular actions of naturally occurring amines in the dog, J. Pharmacol. Exp. Ther. 127, 212–218, 1959. Goldberg, L.I., Dopamine — clinical uses of an endogenous catecholamine, N. Engl. J. Med. 291, 707–710, 1974. Gonzalez, C., Almaraz, L., Obeso, A. and Rigual, R., Carotid body chemoreceptors: from natural stimuli to sensory discharges, Physiol. Rev. 74, 829–898, 1994. Haji, A., Takeda, R. and Okazaki, M., Neuropharmacology of control of respiratory rhythm and pattern in mature mammals, Pharmacol. Ther. 86, 277–304, 2000. Fidone, S.J., Gonzalez, C. and Yoshizaki, K., Putative neurotransmitters in the carotid body: the case for dopamine, Fed. Proc. 39, 2636–2640, 1980. Lahiri, S. and Nishino, T., Inhibitory and excitatory effects of dopamine on carotid chemoreceptors, Neurosci. Lett. 20, 313–318, 1980. Zapata, P., Effects of dopamine on carotid chemoreceptors and baroreceptors in vitro, J. Physiol. (Lond.) 244, 235–251, 1975. Bisgard, G.E., Forster, H.V., Klein, J.P., Manohar, M. and Bullard, V.A., Depression of ventilation by dopamine in goats — effects of carotid body excision, Respir. Physiol. 41, 379–392, 1980. Black, A.M., Comroe, J.H. Jr. and Jacobs, L., Species difference in carotid body response of cat and dog to dopamine and serotonin, Am. J. Physiol. 223, 1097–1102, 1972. Welsh, M., Heistad, D. and Abboud, F., Depression of ventilation by dopamine in man, J. Clin. Invest. 61, 708–713, 1978. Ide, T., Shirahata, M., Chou, C.L. and Fitzgerald, R.S., Effects of a continuous infusion of dopamine on the ventilatory and carotid body responses to hypoxia in cats, Clin. Exp. Pharmacol. Physiol. 22, 658–664, 1995. Loos, N., Haouzi, P. and Marchal, F., Mechanisms of ventilatory inhibition by exogenous dopamine in cats, J. Appl. Physiol. 84, 1131–1137, 1998. Huckauf, H., Ramdohr, B. and Schroder, R., Dopamine induced hypoxemia in patients with left heart failure, Int. J. Clin. Pharmacol. Biopharm. 14, 217–224, 1976. van de Borne, P., Oren, R. and Somers, V.K., Dopamine depresses minute ventilation in patients with heart failure, Circulation 98, 126–131, 1998. Ward, D.S. and Bellville, J.W., Reduction of hypoxic ventilatory drive by dopamine, Anesth. Analg. 61, 333–337, 1982. Lahiri, S., Nishino, T., Mokashi, A. and Mulligan, E., Interaction of dopamine and haloperidol with O2 and CO2 chemoreception in carotid body, J. Appl. Physiol. 49, 45–51, 1980. Ward, D.S. and Bellville, J.W., Effect of intravenous dopamine on hypercapnic ventilatory response in human, J. Appl. Physiol. 55, 1418–1425, 1983. Sabol, S.J. and Ward, D.S., Effect of dopamine on hypoxic–hypercapnic interaction in humans, Anesth. Analg. 66, 619–624, 1987. Nishino, T. and Lahiri, S., Effects of dopamine on chemoreflexes in breathing, J. Appl. Physiol. 50, 892–897, 1981. Cherniack, N.S., von Euler, C., Homma, I. and Kao, F.F., Experimentally induced Cheyne–Stokes breathing, Respir. Physiol. 37, 185–200, 1979.
808 43. 44. 45.
46. 47.
48.
49.
50.
51.
52.
53. 54.
55.
56.
57.
58.
Ward and Karan Naughton, M.T., Pathophysiology and treatment of Cheyne–Stokes respiration, Thorax 53, 514–518, 1998. Lahiri, S., Hsiao, C., Zhang, R., Mokashi, A. and Nishino, T., Peripheral chemoreceptors in respiratory oscillations, J. Appl. Physiol. 58, 1901–1908, 1985. Osanai, S., Akiba, Y., Fujiuchi, S., Nakano, H., Matsumoto, H., Ohsaki, Y. and Kikuchi, K., Depression of peripheral chemosensitivity by a dopaminergic mechanism in patients with obstructive sleep apnoea syndrome, Eur. Respir. J. 13, 418–423, 1999. Johnson, R.L., Low-dose dopamine and oxygen transport by the lung, Circulation 98, 7–99, 1998. O’Halloran, K.D., Herman, J.K. and Bisgard, G.E., Respiratory-related pharyngeal constrictor muscle activity in awake goats, Respir. Physiol. 116, 9–23, 1999. Cabezas, G.A., Lezama, Y., Vidal, A. and Velasco, M., Inhaled dopamine induces bronchodilatation in patients with bronchial asthma, Int. J. Clin. Pharmacol. Ther. 37, 510–513, 1999. Lin, Y.S., Gu, Q. and Lee, L.Y., Activation of dopamine D2-like receptors attenuates pulmonary C-fiber hypersensitivity in rats, Am. J. Respir. Crit. Care Med. 167, 1096–1101, 2003. Murphy, M.B., Murray, C. and Shorten, G.D., Fenoldopam: a selective peripheral dopamine-receptor agonist for the treatment of severe hypertension, N. Engl. J. Med. 345, 1548–1557, 2001. Azzam, Z.S., Saldias, F.J., Comellas, A., Ridge, K.M., Rutschman, D.H. and Sznajder, J.I., Catecholamines increase lung edema clearance in rats with increased left atrial pressure, J. Appl. Physiol. 90, 1088–1094, 2001. Drolet, B., Zhang, S., Deschenes, D., Rail, J., Nadeau, S., Zhou, Z., January, C.T. and Turgeon, J., Droperidol lengthens cardiac repolarization due to block of the rapid component of the delayed rectifier potassium current, J. Cardiovasc. Electrophysiol. 10, 1597–1604, 1999. Ward, D.S., Stimulation of hypoxic ventilatory drive by droperidol, Anesth. Analg. 63, 106–110, 1984. Bascom, D.A., Clement, I.D., Dorrington, K.L. and Robbins, P.A., Effects of dopamine and domperidone on ventilation during isocapnic hypoxia in humans, Respir. Physiol. 85, 319–328, 1991. Aho, M., Erkola, O., Kallio, A., Scheinin, H. and Korttila, K., Dexmedetomidine infusion for maintenance of anesthesia in patients undergoing abdominal hysterectomy, Anesth. Analg. 75, 940–946, 1992. Belleville, J.P., Ward, D.S., Bloor, B.C. and Maze, M., Effects of intravenous dexmedetomidine in humans I. Sedation, ventilation and metabolic rate, Anesthesiology 77, 1125–1133, 1992. Oates, J.A. and Brown, N.J., Antihypertensive agents and the drug therapy of hypertension, in The Pharmacological Basis of Therapeutics, Hardman, J.G. and Limbird, L.E., eds., New York, McGraw-Hill, pp. 871–900, 2001. Unnerstall, J.R., Kopajtic, T.A. and Kuhar, M.J., Distribution of a2-agonist binding sites in the rat and human central nervous system: analysis of some functional, anatomic correlates of the pharamcologic effects of clonidine and related adrenergic agents, Brain Res. Rev. 7, 69–101, 1984.
Cardiovascular Drugs and the Control of Breathing 59.
60. 61.
62. 63.
64.
65.
66.
67.
68. 69. 70.
71. 72. 73.
74. 75.
76.
809
O’Halloran, K.D., Herman, J.K. and Bisgard, G.E., Ventilatory effects of alpha(2)-adrenoceptor blockade in awake goats, Respir. Physiol. 126, 29–41, 2001. Benhamou, D., Veillette, Y., Narchi, P. and Ecoffey, C., Ventilatory effects of premedication with clonidine, Anesth. Analg. 73, 799–803, 1991. Narchi, P., Benhamou, D., Hamza, J. and Bouaziz, H., Ventilatory effects of epidural clonidine during the first 3 hours after caesarean section, Acta Anaesthesiol. Scand. 36, 791–795, 1992. Ooi, R., Pattison, J. and Feldman, S.A., The effects of intravenous clonidine on ventilation, Anaesthesia 46, 632–633, 1991. Bisgard, G.E., Mitchell, R.A. and Herbert, D.A., Effects of dopamine, norepinephrine and 5-hydroxytryptamine on the carotid body of the dog, Respir. Physiol. 37, 61–80, 1979. McCrimmon, D.R. and Lalley, P.M., Inhibition of respiratory neural discharges by clonidine and 5-hydroxytryptophan, J. Pharmacol. Exp. Ther. 222, 771–777, 1982. Coles, S.K., Ernsberger, P. and Dick, T.E., Post-hypoxic frequency decline does not depend on alpha2-adrenergic receptors in the adult rat, Brain Res. 794, 267–273, 1998. Ernsberger, P., Kou, Y.R. and Prabhakar, N.R., Carotid body I1-imidazoline receptors: binding, visualization and modulatory function, Respir. Physiol. 112, 239–251, 1998. Nguyen, D., Abdul-Rasool, I., Ward, D.S., Hsieh, J., Kobayashi, D., Hadlock, S., Singer, F. and Bloor, B., Ventilatory effects of dexmedetomidine, atipamezole, and isoflurane in dogs, Anesthesiology 76, 573–579, 1992. Zornow, M.H., Ventilatory, hemodynamic and sedative effects of the a2 adrenergic agonist, dexmedetomidine, Neuropharmacology 30, 1065–1071, 1991. Ward, D.S., Respiratory effects of a2-adrenoceptor agonists, Anaesth. Pharm. Rev. 1, 263–267, 1993. Hedrick, M.S., Ryan, M.L. and Bisgard, G.E., Recurrent laryngeal nerve activation by alpha(2) adrenergic agonists in goats, Respir. Physiol. 101, 129– 137, 1995. Folgering, H., Central beta-adrenergic effects on the control of ventilation in cats, Respiration 39, 131–138, 1980. Eldridge, F.L. and Gill-Kumar, P., Mechanisms of hyperpnea induced by isoproterenol, Respir. Physiol. 40, 349–363, 1980. Heistad, D., Wheeler, R., Mark, A., Schmid, P. and Abboud, F.M., Effects of adrenergic stimulation on ventilation in man, J. Clin. Invest. 51, 1369–1475, 1972. Folgering, H., Ponte, J. and Sadig, T., Adrenergic mechanisms and chemoreception in the carotid body of the cat and rabbit, J. Physiol. (Lond.) 325, 1–21, 1982. Yu, B.H., Mills, P.J., Ziegler, M.G. and Dimsdale, J.E., Sympathetic and respiratory responses to hypoxia in essential hypertension, Clin. Exp. Hypertens. 21, 249–262, 1999. Clar, C., Dorrington, K.L. and Robbins, P.A., Ventilatory effects of 8 h of isocapnic hypoxia with and without beta-blockade in humans, J. Appl. Physiol. 86, 1897–1904, 1999.
810 77.
78.
79.
80.
81. 82. 83.
84.
85.
86.
87.
88. 89.
90.
91.
92.
Ward and Karan Clar, C., Dorrington, K.L., Fatemian, M. and Robbins, P.A., Cardiovascular effects of 8 h of isocapnic hypoxia with and without beta-blockade in humans, Exp. Physiol. 85, 557–565, 2000. Freund, B.J., Claybaugh, J.R., Dice, M.S. and Hashiro, G.M., Hormonal and vascular fluid responses to maximal exercise in trained and untrained males, J. Appl. Physiol. 63, 669–675, 1987. Ohtake, P.J., Walker, J.K. and Jennings, D.B., Renin-angiotensin system stimulates respiration during acute hypotension but not during hypercapnia, J. Appl. Physiol. 74, 1220–1228, 1993. Jennings, D.B., Walker, J.K. and Ohtake, P.J., Central effects and interactions of the renin-angiotensin system and vasopressin on ventilation and PaCO2, in Ventral Brainstem Mechanisms and Control of Respiration and Blood Pressure, Trouth, C.O., Millis, R.M., Kiwull-Schone, H. and Schlafke, M., eds., New York, Marcel Dekker, pp. 203–224, 1995. Lam, S.Y. and Leung, P.S., A locally generated angiotensin system in rat carotid body, Regul. Pept. 107, 97–103, 2002. Allen, A.M., Angiotensin AT1 receptor-mediated excitation of rat carotid body chemoreceptor afferent activity, J. Physiol. (Lond.) 510, 773–781, 1998. Leung, P.S., Lam, S.Y. and Fung, M.L., Chronic hypoxia upregulates the expression and function of AT(1) receptor in rat carotid body, J. Endocrinol. 167, 517–524, 2000. Kc, P., Haxhiu, M.A., Tolentino-Silva, F.P., Wu, M., Trouth, C.O. and Mack, S.O., Paraventricular vasopressin-containing neurons project to brain stem and spinal cord respiratory-related sites, Respir. Physiol. Neurobiol. 133, 75–88, 2002. Ferguson, A.V., Beckmann, L.M. and Fisher, J.T., Effects of subfornical organ stimulation on respiration in the anesthetized rat, Can. J. Physiol. Pharmacol. 67, 1097–1101, 1989. Walker, J.K. and Jennings, D.B., Angiotensin mediates stimulation of ventilation after vasopressin V1 receptor blockade, J. Appl. Physiol. 76, 2517–2526, 1994. Walker, J.K. and Jennings, D.B., During acute hypercapnia vasopressin inhibits an angiotensin drive to ventilation in conscious dogs, J. Appl. Physiol. 79, 786–794, 1995. Jennings, D.B., Cardiorespiratory effects of prolonged angiotensin II block in resting conscious dogs, Can. J. Physiol. Pharmacol. 79, 825–830, 2001. Walker, J.K. and Jennings, D.B., Respiratory effects of pressor and depressor agents in conscious rats, Can. J. Physiol. Pharmacol. 76, 707–714, 1998. Ohtake, P.J. and Jennings, D.B., Angiotensin II stimulates respiration in awake dogs and antagonizes baroreceptor inhibition, Respir. Physiol. 91, 335–351, 1993. Heitman, S.J. and Jennings, D.B., Angiotensin II modulates respiratory and acid–base responses to prolonged hypoxia in conscious dogs, Am. J. Physiol. 275, R390–R399, 1998. Jennings, D.B. and Lockett, H.J., Angiotensin stimulates respiration in spontaneously hypertensive rats, Am. J. Physiol. 278, R1125–R1133, 2000.
Cardiovascular Drugs and the Control of Breathing 93.
94.
95.
96.
97.
98. 99.
100.
101.
102.
103.
104.
105.
106.
811
Montgomery, H.E., Marshall, R., Hemingway, H., Myerson, S., Clarkson, P., Dollery, C., Hayward, M., Holliman, D.E., Jubb, M., World, M., Thomas, E.L., Brynes, A.E., Saeed, N., Barnard, M., Bell, J.D., Prasad, K., Rayson, M., Talmud, P.J. and Humphries, S.E., Human gene for physical performance, Nature 393, 221–222, 1998. Gayagay, G., Yu, B., Hambly, B., Boston, T., Hahn, A., Celermajer, D.S. and Trent, R.J., Elite endurance athletes and the ACE I allele — the role of genes in athletic performance, Hum. Genet. 103, 48–50, 1998. Woods, D.R., Pollard, A.J., Collier, D.J., Jamshidi, Y., Vassiliou, V., Hawe, E., Humphries, S.E. and Montgomery, H.E., Insertion/deletion polymorphism of the angiotensin I-converting enzyme gene and arterial oxygen saturation at high altitude, Am. J. Respir. Crit. Care Med. 166, 362–366, 2002. Fisher, T.E. and Bourque, C.W., The function of Ca2þ channel subtypes in exocytotic secretion: new perspectives from synaptic and non-synaptic release, Prog. Biophys. Mol. Biol. 77, 269–303, 2001. Lopez-Barneo, J., Pardal, R., Montoro, R.J., Smani, T., Garcia-Hirschfeld, J. and Urena, J., Kþ and Ca2þ channel activity and cytosolic [Ca2þ] in oxygensensing tissues Respir. Physiol. 115, 215–227, 1999. Hackett, P. and Roach, R., Medical therapy of altitude illness, Ann. Emerg. Med. 16, 980–986, 1987. Overholt, J.L. and Prabhakar, N.R., Ca2þ current in rabbit carotid body glomus cells is conducted by multiple types of high-voltage-activated Ca2þ channels, J. Neurophysiol. 78, 2467–2474, 1997. Obeso, A., Rocher, A., Fidone, S. and Gonzalez, C., The role of dihydropyridine-sensitive Ca2þ channels in stimulus-evoked catecholamine release from chemoreceptor cells of the carotid body, Neuroscience 47, 463–472, 1992. Chapman, R.W., Effect of verapamil on ventilation and chemical control of breathing in anesthetized rats, Can. J. Physiol. Pharmacol. 63, 1608–1611, 1985. Long, G.R., Filuk, R., Balakumar, M., Easton, P.A. and Anthonisen, N.R., Ventilatory response to sustained hypoxia: effect of methysergide and verapamil, Respir. Physiol. 75, 173–182, 1989. Ohdaira, T., Kobayashi, T., Tanaka, M., Chowdhury, M.F., Ahn, B., Masuda, A., Sakakibara, Y. and Honda, Y., Effect of verapamil on ventilatory and circulatory responses to hypoxia and hypercapnia in normal subjects, Jpn. J. Physiol. 42, 765–777, 1992. Carta, F., Bianchi, M., Argenton, S., Cervi, D., Marolla, G., Tamburini, M., Breda, M., Fantoni, A. and Panerai, A.E., Effect of nifedipine on morphineinduced analgesia, Anesth. Analg. 70, 493–498, 1990. Ruiz, F., Dierssen, M., Florez, J. and Hurle, M.A., Potentiation of acute opioid-induced respiratory depression and reversal of tolerance by the calcium antagonist nimodipine in awake rats, Naunyn Schmiedebergs Arch. Pharmacol. 348, 633–637, 1993. Kishioka, S., Ko, M.C. and Woods, J.H., Diltiazem enhances the analgesic but not the respiratory depressant effects of morphine in rhesus monkeys, Eur. J. Pharmacol. 397, 85–92, 2000.
812
Ward and Karan
107. Javaheri, S. and Guerra, L., Lung function, hypoxic and hypercapnic ventilatory responses, and respiratory muscle strength in normal subjects taking oral theophylline, Thorax 45, 743–747, 1990. 108. Watt, A.H. and Routledge, P.A., Adenosine stimulates respiration in man, Br. J. Clin. Pharmacol. 20, 503–506, 1985. 109. Maxwell, D.L., Fuller, R.W., Nolop, K.B., Dixon, C.M.S. and Hughes, J.M.B., Effects of adenosine on ventilatory responses to hypoxia and hypercapnia in humans, J. Appl. Physiol. 61, 1762–1766, 1986. 110. Yamamoto, M., Nishimura, M., Kobayashi, S., Akiyama, Y., Miyamoto, K. and Kawakami, Y., Role of endogenous adenosine in hypoxic ventilatory response in humans: a study with dipyridamole, J. Appl. Physiol. 76, 196–203, 1994. 111. Easton, P.A. and Anthonisen, N.R., Ventilatory response to sustained hypoxia after pretreatment with aminophylline, J. Appl. Physiol. 64, 1445–1450, 1988. 112. Georgopoulos, D., Holtby, S.G., Berezanski, D. and Anthonisen, N.R., Aminophylline effects on ventilatory response to hypoxia and hyperoxia in normal adults, J. Appl. Physiol. 67, 1150–1156, 1989. 113. Easton, P.A., Slykerman, L.J. and Anthonisen, N.R., Ventilatory response to sustained hypoxia in normal adults, J. Appl. Physiol. 61, 906–911, 1986. 114. Gourine, A.V., Llaudet, E., Thomas, T., Dale, N. and Spyer, K.M., Adenosine release in nucleus tractus solitarii does not appear to mediate hypoxia-induced respiratory depression in rats, J. Physiol. (Lond.) 544, 161–170, 2002. 115. Javaheri, S., Parker, T.J., Wexler, L., Liming, J.D., Lindower, P. and Roselle, G.A., Effect of theophylline on sleep-disordered breathing in heart failure, N. Engl. J. Med. 335, 562–567, 1996. 116. Maxwell, D.L., Fuller, R.W., Dixon, C.M.S., Cuss, F.M.C. and Barnes, P.J., Ventilatory effects of substance P, vasoactive intestinal peptide, and nitroprusside in humans, J. Appl. Physiol. 68, 295–301, 1990. 117. Prabhakar, N.R., NO and CO as second messengers in oxygen sensing in the carotid body, Respir. Physiol. 115, 161–168, 1999. 118. Chugh, D.K., Katayama, M., Mokashi, A., Bebout, D.E., Ray, D.K. and Lahiri, S., Nitric oxide-related inhibition of carotid chemosensory nerve activity in the cat, Respir. Physiol. 97, 147–156, 1994. 119. Nakano, H., Lee, S.D., Ray, A.D., Krasney, J.A. and Farkas, G.A., Role of nitric oxide in thermoregulation and hypoxic ventilatory response in obese Zucker rats, Am. J. Respir. Crit. Care Med. 164, 437–442, 2001. 120. Prabhakar, N.R., Kumar, G.K., Chang, C.H., Agani, F.H. and Haxhiu, M.A., Nitric oxide in the sensory function of the carotid body, Brain Res. 625, 16–22, 1993. 121. Summers, B.A., Overholt, J.L. and Prabhakar, N.R., Nitric oxide inhibits L-type Ca2þ current in glomus cells of the rabbit carotid body via a cGMP-independent mechanism, J. Neurophysiol. 81, 1449–1457, 1999. 122. Iturriaga, R., Villanueva, S. and Mosqueira, M., Dual effects of nitric oxide on cat carotid body chemoreception, J. Appl. Physiol. 89, 1005–1012, 2000.
Cardiovascular Drugs and the Control of Breathing
813
123. Iturriaga, R., Alcayaga, J. and Rey, S., Sodium nitroprusside blocks the cat carotid chemosensory inhibition induced by dopamine, but not that by hyperoxia, Brain Res. 799, 26–34, 1998. 124. Lipton, A.J., Johnson, M.A., Macdonald, T., Lieberman, M.W., Gozal, D. and Gaston, B., S-nitrosothiols signal the ventilatory response to hypoxia, Nature 413, 171–174, 2001. 125. McQueen, D.S. and Ribeiro, J.A., Effects of ouabain on carotid body chemoreceptor activity in the cat, J. Physiol. (Lond.) 335, 221–235, 1983. 126. Schobel, H.P., Ferguson, D.W., Clary, M.P. and Somers, V.K., Differential effects of digitalis on chemoreflex responses in humans, Hypertension 23, 302–307, 1994.
AUTHOR INDEX
Ahshuler, K.Z., 760, 775 Akiyama, Y., 552, 569 Alexander, C.M., 521, 525, 538, 558, 566 Allen, G.D., 529, 564 Allsop, P., 532, 565 Amagasa, S., 783, 791 Amin, H.M., 550, 569 Andres, L.P., 303, 311–312 Andriano, K.P., 550, 561 Andronikou, S., 657, 680 Ansved, T., 330, 352 Aqleh, K.A., 548, 568 Arunasalam, K., 545, 567 Audenaert, S.M., 528, 563 Avramov, M.N., 557, 570 Axen, K., 83, 99
Babenco, H.D., 531–532, 535, 538, 550, 565–566, 569 Backman, L., 391, 415 Badrinath, S., 557, 570 Bailey, P.L., 469, 477, 496–497, 500, 506, 509, 544, 550, 554, 561 Baker R.W., 390, 396, 415 T.L., 173 Bali, I.M., 565 Ballantyne D., 44, 64 J.C., 459, 461, 485, 491
Barnard, J.D., 459, 502, 512 Barzilay, E., 537, 566 Beattie C., 487, 511 W.S., 437, 494 Becker, L.D., 555, 570, 751, 769 Bell, G.D., 526, 562 Belleville, J., 477, 509, 517, 527, 548, 563, 568 Benhamou, D., 478, 509 Berkenbosch, A., 115, 129 Berthon-Jones, M., 404, 410, 419 Bickler, P.E., 529, 564 Bisgard, G.E., 161, 167, 200–201, 204, 216, 218, 799, 809 Bloor, B.C., 477, 509 Blouin, R.T., 531–532, 535, 550, 565 Bolser, D.C., 220 Bolton, D.P., 761, 776 Borgbjerg, F.M., 439, 495 Bouillon, T., 117, 124, 550, 561 Boulet, L.-P., 247, 257 Bourke, D.L., 113, 128, 520, 527, 536, 553, 556–557, 565, 577 Bowen, D.J., 545, 567 Bradley S.R., 24–25, 57 T.D., 399, 417 Bragg, P., 109, 127 Brickleyk, S.G., 595, 644 Brodner, G., 436, 493 Bromage, P.R., 466, 505
815
816 Brouillette, R.T., 540, 548, 566 Brown C.R., 527, 563 J.T., 465, 505 Bryan, A.C., 703, 729 Burki, N.K., 390, 396, 415 Buyse, B., 396, 416 Cabello, J., 412, 422 Caldwell, C.B., 455, 501 Campbell, D., 546, 566 Camporesi, E.M., 466, 505 Cantineau, J.P., 437, 464, 493, 753, 771 Carlisle, C.C., 401, 418 Carr, D.B., 459, 461, 485, 491 Carter, A.S., 526, 562 Cegla, U.H., 517, 560 Celli, B.R., 410, 422 Cepeda, M.S., 545, 567 Chang, F., 744, 767 Chapman, K.R., 396, 417 Chen E., 399, 410, 417 Z., 24, 37, 56 Cheng, G., 593, 642 Cheong, T.H., 516–517, 521, 560 Cherniak, N.S., 375, 382 Choi, S.D., 527–528, 561, 564 Clark F.J., 611, 646 L.D., 557, 570 Clergue, F., 455, 501 Cohen R., 527, 563 S.E., 478, 509 Coles, S.K., 162, 201 Connelly, C.A., 272, 288 Conwell, D.L., 558, 562 Coombs, J.H.B., 555, 569 Corris, P.A., 517, 560 Cote, C.J., 528, 563 Cowen, E.R., 389, 392, 409, 415 Crawford, R.D., 368, 378 Croci, M., 388–389, 415 Cronin, A., 761, 776 Cui, N., 45, 65 Cull-Candy, S.G., 595, 644 Dahan, A., 105, 130, 439, 495, 497, 524, 545, 561, 567, 574, 632, 750, 752, 770, 802 Daniels, A.L., 528, 563
Author Index Davenport H.T., 545, 567 P.W., 192, 220 Davies D.G., 25, 57 H., 484, 491 J.A., 239, 255 R.J., 113, 128 R.O., 668, 683 Daykin, A.P., 545, 567 Dean, J.B., 44, 64 de Backer, W., 398–399, 417 deGoede, J., 115, 129 deGroot, W.J., 314, 338 Dejours, P., 105 Dekin, M.S., 160, 206 Delavault, E., 537, 566 de Lorenzo, A., 388, 415 Delpierre, S., 516–517, 559 de Miguel, J., 412, 422 Dempsey, J.A., 167, 173, 208, 365, 372, 376–377 Denavit-Saubie, 595, 644 D’Honneur, G., 788 Dick, T.E., 162, 201 Dirkes, W.E., 756–757, 773 Distenfeld, S., 83, 99 Dogas, Z., 668, 683–684 Doi, K., 583, 636 Dong, X.W., 172, 211 Douglas, R.M., 37, 61 Downes, J.J., 546, 568 Downie, D.L., 594, 643, 790 Drummond, G.B., 531, 564, 711, 724, 728, 730 Dudziak, R., 453, 500 Dueck, R., 529, 564 Duffin, J., 113, 127, 531, 564 Duncan, S.R., 477, 509 Dundee, J.W., 459, 502, 565 Dureuil, B., 753, 771 Durrani, Z., 453, 500 Dutton, R.E., 25, 57 Dwinell, M.R., 366, 378
Eastwood, P.R., 711–712, 714 Ecoffey, C., 478, 509, 557, 565, 577, 633 Edwards, M.W., 668, 683 Eisenach, J., 478, 508–509 El-Bohy, A.A., 196, 223 Eldridge, F.L., 160, 203, 206–207, 420 Emirgil, C., 409, 415
Author Index Enson, Y., 391, 409, 416 Eriksson, L.I., 742, 766 Escarraga, L.A., 744, 767 Esposito, B.F., 550, 569 Farrant, M., 595, 644 Farrar, J.T., 545, 567 Fatemian, M., 171, 211 Feldman, S.A., 477, 509 Fencl, V., 47, 56 Fink, B.R., 84, 99 Finn, H., 527, 563 Fitzgerald, R.S., 781 Fleury, B., 579, 634 Flynn, K., 555–556, 570 Foerster, O., 75, 93 Ford, E.S., 384, 412 Forrest, W.H., Jr, 527, 563 Forster A., 161, 167, 521, 524, 526, 536, 560–561, 563, 565 H.V., 167, 173, 208 Foschino Barbaro, M.P., 407, 420 Foutz, A.S., 595, 644 Franks, J., 439, 495 Friberg, D., 330, 352 Froese, A.B., 703, 729 Fuller D.D., 189, 191, 196 R.W., 803–804, 812 Gal, T.J., 786, 790 Galland, B.C., 761, 776 Gamsu, G., 754, 771 Gan, T.J., 453, 500 Garay, S.M., 399, 417 Garstka, G., 117, 124, 550, 561 Gasser, J.C., 517, 527, 563 Gautier, H., 579–580, 634 Gelb, A.W., 632, 751, 758 Gemperle, M., 536, 565 Gerhardstein, D.C., 241, 256 Germain, M., 531, 564 Gesell, R., 527, 563 Gilles, D.M.M., 531, 564 Gilmartin, J.J., 517, 560 Ginsberg, B., 453, 500 Glass, P.S.A., 550, 552, 569 Gleed, R.D., 183, 217 Glynn, C.J., 459, 502, 512 Goetzinger, C.P., 83, 99 Gold, A.R., 398, 417
817 Golder, F.J., 189, 192–193, 220 Goldring, R.M., 517, 560 Gonzalez, C., 796, 801 Goodman, N.W., 532, 564–565 Gorcs, T.J., 24, 56 Gourine, A.V., 804, 812 Gozal D., 162, 171–173, 204, 207, 209–211 E., 162, 204, 207, 209 Grant, I.S., 529, 532, 564 Grassino, A.E., 390, 416 Greenberg, H.E., 517, 560 Gross, J.B., 455, 501, 521, 526–527, 557–558, 561–562, 565, 570, 577, 633 Grounds, R.M., 531–532, 564–565 Gust A., 450, 499 R., 450, 499 Guyenet, P.G., 162, 204
Haas, F., 83, 99 Haji, A., 594, 635 Hamilton, R.D., 238, 255 Hammer, J., 536, 565 Hamunen, K., 544, 548, 567 Hamza, J., 557, 565, 577, 633 Han, F., 399, 410, 417 Hanacek, J.A., 239, 255 Hansdottir, V., 466, 506–507 Hansen, E.T., 527, 563 Haouzi, P., 796, 807 Harf, A., 389, 416 Harper M.H., 548, 555, 568 R.M., 311 Harrer, G., 83, 99 Hasegawa, R., 748, 769 Hayden, J., 527, 563 Hedenstierna, G., 471, 504 Hedrick, M.S., 799, 809 Hemmings, H.C., 585, 588–589, 637–638 Henry, J.P., 388, 415 Hensley, M.J., 555, 570 Hershenson, M., 540, 548, 566 Hertle, L., 436, 493 Heymans, C.J.F., 3, 781, 790 Heywood, P., 52, 69 Hickey, R.F., 548, 555, 568 Higenbottam, T., 231, 253 Hill, N.S., 271, 288 Himal, H.S., 396, 417 Hirshman, C.A., 528, 563
818 Holte, K., 486, 491 Holter, P.H., 391, 416 Honore, E., 591, 631 Hornbein, T.F., 365, 372, 376–377 Horner, R.L., 238, 255, 389, 416, 418 Hudson, L.D., 517, 560 Huey, K.A., 168, 209, 366, 378 Igarashi, A., 783, 791 Irwin, T.S., 247, 257 Iseline-Chaves, I.A., 550, 569 Isono, S., 249, 259, 739, 746–747, 764–765 Jackson C., 240, 256 M., 231, 253 James, D., 37, 61 Jammes, Y., 516–517, 559 Jamrozik, K., 436, 485–486, 491 Jarvis, D.A., 477, 509 Javaheri, S., 299, 398–399, 405, 407, 417, 419–420, 804, 812 Jensen, R.L., 390, 416 Jhaveri, R.M., 552, 569 Jiang, C., 45, 65 Johansson, H., 471, 504 Johnson, S.M., 156, 199 Jones, R.L., 397, 417 Jonsson, M., 783 Jorgensen, N.K., 50, 65 Kafer, E.R., 465, 505 Kasaba, T., 583, 636 Katz, J., 459–460, 491, 503 Kaufman, R.D., 517, 527, 548, 563, 568 Kavanagh, B.P., 459–460, 491, 503 Kavey, N.B., 760, 775 Kay, B., 528, 563 Keenan, D.J., 459, 502 Kehlet, H., 450, 486, 491, 493–494, 756–757, 773 Keifer, J.C., 761, 776 Kelly, T.M., 390, 416 Kemp, R.A., 546, 568 Kendig, J.J., 593, 642 Kennedy, W.F., Jr., 529, 564 Khan, A.U., 552, 569 Khoo, M.C.K., 106, 125 Kille, J.F., 300, 309 Kimura, H., 396, 417 Kinkead, R., 186 Kirvela, O.A., 109, 127, 166
Author Index Kleeberger, S., 290 Kline, D.D., 160, 166, 169, 202–203, 206 Knill, R., 529, 531, 564, 566, 632, 751, 758–760, 773, 775 Knuth, S.L., 517, 560 Kochi, T., 752, 770 Koenig, J.S., 324, 349 Kosaka, Y., 583, 636 Koshiya, N., 162, 204 Koury, S.I., 537, 565 Kozar, L., 240, 256 Kress, J.P., 391, 416 Kuna, S.T., 332, 350–351, 353–354 Kunitomo, F., 396, 417
Labaille, T., 455, 501 Lahiri, S., 375, 382, 668, 683, 797 Lakshminarayan, S., 517, 560 Lambertsen, C.J., 546, 568 Lang, S., 231, 252 Larkin, E.K., 400, 418 Larsson, L.E., 528, 563 Latasch, L., 453, 500 Lau, M., 109, 127 Launois, S., 579, 634 Lavie, P., 408, 421 Lee B., 513, 559 K.K., 486, 511 L.-Y., 241, 256 Leech, J.A., 392, 407, 417 Lees, M.H., 538, 566 Lehmann, K.A., 537, 554, 566 Leino, K., 557, 570 Leiter, J.C., 39, 62, 517, 560 Leusen, I.R., 21 Lin, C.C., 399, 410, 417 Lindsey, B.G., 166, 207 Linton, R.A., 113, 128, 795 Liposits, Z., 24, 56 Lister, R.E., 546, 566 Litman, R.S., 521, 536, 562, 565–566 Llaudet, E., 804, 812 Lloyd, J.W., 459, 502, 512 Loeschcke, H.H., 22, 66, 300, 309 Loesche, H.H., 22, 367, 378 Looi-Lyons, L.C., 445, 449 Loos, N., 796, 807 Lopata, M., 390, 396–397, 399, 416 Lopez-Barneo, J., 6, 17, 801, 811 Lopez-Lopez, J.R., 6, 17 Lu, J.K., 471, 506–507
Author Index Lue, F., 399, 417 Lundh, R., 471, 504 Lynn, A.M., 545, 567
Mackenzie, N., 529, 532, 564 Mahan, R.A., 556, 570 Mainka, F., 537, 554, 566 Maiolo, C., 388, 415 Mak, K.H., 516–517, 521, 560 Malit, L.A., 556, 565 Man, G.C., 397, 417 Manikian, B., 437, 464, 493 Mann, C.B., 105, 130 Marc, I., 517, 560 Marchal, F., 796, 807 Markous, N., 396, 416 Marx, C.M., 556, 570 Masa, J.F., 410, 422 Maxwell, D.L., 531, 564, 803–804, 812 McCarthy, R.J., 483, 494 McCrimmon, D.R., 160, 206 McCullough, R.E., 528, 544, 563, 566 McKeating, K., 565 McLeod, G., 484, 491 McNicol, G.W., 546, 566 McQueen, D.S., 805, 809, 813 Mezzanotte, W.S., 325, 350 Mikkelsen, S., 456, 501 Mildh, L., 109, 127, 166, 557, 570 Milhorn, D.E., 166 Miller, T.B., 47, 56 Millhorn, D.E., 160, 203, 206–207, 420 Millman, R.P., 401, 418 Milsom, W.K., 129, 167–168, 202 Mitchell G.S., 129, 156, 160, 167–168, 189, 191, 196, 199, 202, 206 R.A., 22, 296, 305, 367, 378 R.K., 537, 565 Mohan, R.M., 113, 127 Moiniche, S., 456, 501 Mokashi, A., 657, 680 Mokdad, A.H., 384, 412 Monge, C.C., 375, 382 Monheim, L.M., 513–514, 559 Montravers, P., 526 Moorjani, N., 459, 503 Moote, C.A., 709, 759–760, 773, 775 Mora, C.T., 520, 525–526, 561–562 Morales, P., 459, 502 Morel, D.R., 536, 565 Morrell, M.J., 52, 69
819 Morris, K.F., 166, 173, 200, 202, 207 Morrow, J.B., 558, 562 Mortero, R.F., 557, 570 Mortola, J.P., 183–185, 217 Mottram, S.D., 754, 771 Muir, A.D., 756, 774 Muller-JSuur, R., 391, 415 Mulroy, M.F., 555, 569 Murray, A, 527, 563 Nagyova, B., 535 Narchi, P., 478, 509 National Institutes of Health, 385 Nattie, E.E., 368 Nespeca, M.K., 545, 567 Neubauer, J.A., 167 Nielsen C.H., 466, 505 K., 439, 495 Nieuwenhuijs, D., 532, 535, 565 Nilsson, K., 528, 563 Nishimura, M., 552, 569, 804, 812 Nishino, T., 710, 748, 752, 769–770 Norris, E.J., 487, 511 Nottingham, S., 39, 62 Nurse, C.A., 653 Offenstadt, G., 579–580, 634 O’Halloran, K.D., 798 Okada, Y., 24, 37, 56 Okazaki, M., 594, 635 Olievier, I.C.W., 574, 632 Oliven, A., 330 Olofsen, E., 802 Olsen, G.D., 538, 566 Olson, L.G., 555, 570 Onal, E., 390, 392, 396–397, 399, 407, 416–417 Ooi, R., 477, 509 Orr, I.A., 459, 502 Ortiz, T., 190, 197, 220 Otto-Smith, M.R., 272, 288 Ou, L.C., 271, 288 Overholt, J.L., 160, 166, 203 Paalzow, L., 520–521, 524, 561 Pace, N.L., 554, 561 Palta, M., 399, 414 Pancrazio, J.J., 586, 637 Pappenheimer, J.R., 47, 56 Pardal, R., 801, 811 Park, W.Y., 486, 511
820 Parker, R.I., 556, 570 Pastor, J., 459, 502 Patel, A.J., 591, 631 Paton, J.Y., 302–303, 311 Pattison, J., 477, 509 Paulson, B.A., 546, 555, 568, 570, 751, 769 Pedersen, K.A., 50, 65 Peiser, J., 408, 421 Pelosi, P., 388–389, 415 Peng, Y., 169, 205–206 Penon, C., 478, 509 Peracchia, C., 31, 63–64 Philip-Joet, F., 517, 560 Picket, C.K., 380, 657, 678 Pickett, C.K., 372, 379 Pleuvry, B.J., 553, 569 Polaner, D.M., 528, 563 Ponte, J., 669, 684 Poole, R., 83, 99 Powell, F.L., 129, 167–168, 202, 209, 366, 369, 378 Power, S.J., 520–521, 527 Prabhakar, N.R., 160, 166, 203 Prokocimer, P., 537, 566 Prutow, R.J., 529, 564 Ramamurthy, S., 553, 569 Randall, N.P.C., 553, 569 Raphael, J.H., 754, 771 Rapoport, D.M., 517, 560 Raymond, L.W., 668, 683 Rebuck, A.S., 396, 417 Reeder, M.K., 756, 774 Refsum, H.E., 391, 416 Reier, P.J., 196, 223 Remmers, J.E., 314, 338 Ren, X., 171, 211 Resta, O., 407, 420 Rezzonico, R., 183, 217 Ribeiro, J.A., 805, 809, 813 Richards, F.J., 134, 153 Rigg, J.R., 436, 485–486, 491, 497, 529, 564, 566, 568 Robbins, P.A., 171, 211 Rochester, D.F., 391, 409, 416 Rodgers, A., 427, 491 Rosenberg, J., 756–758, 761, 764, 773 Rosenbluth, A., 190, 197, 220 Rowley, J.A., 323, 356 Sadler, C.L., 669, 684 Safar, P., 744, 767
Author Index Sampson, M.G., 390, 416 Sandler, A.N., 459–460, 466, 491, 503 Sant’Ambrogio, G., 227, 239, 253 Sant’Anna, G.M., 184–185, 217 Santer, L.J., 537, 565 Santiago, T.V., 552, 569 Sargent, C.W., 302–303, 311 Sarton, E., 439, 495, 497, 532, 535, 545, 565, 752, 770 Sato, M., 742, 766 Saunders, N.A., 555, 570 Schafer, M., 450, 499 Schla¨fke, M.E., 22, 66, 300, 309 Schlame, M., 588–589, 637 Schmid, K., 595, 644 Schug, S., 427, 491 Schwab, R., 401 Seanhy, S.K., 388, 415 Serebrovskaya, L.A., 168, 210 Severinghaus, J.W., 296, 305, 367–368, 378, 668, 683, 742, 766 Sharp, J.T., 388, 415 Shea, S.A., 303, 311–312 Sheller, J.R., 545, 567 Shively, J.G., 555–556, 570 Sieker, H.O., 392, 417 Sin, D.D., 397, 417 Sirois, J.E., 586, 637 Skaggs, E.B., 83, 99 Smith C.A., 365, 372, 376–377, 647 G., 753, 771 J.A., 537, 565 J.C., 598, 636, 645 L.R., 552, 569 T.C., 786, 790 Sobol, B.J., 409, 415 Soroker, D., 537, 566 Spaulding, B.C., 527, 561 Spence, A.A., 753, 771 Spens, H.J., 531, 564 Steen, S.N., 517, 527, 560, 563 Stockwell, M., 231, 252 Stone, C.K., 537, 565 Strohl, K.P., 273, 288, 393, 401, 419 Stuth, E.A.E., 582, 632, 668, 683–684 Sullivan, C.E., 240, 256, 404, 410, 419 Sunzel, M., 520–521, 524, 561 Sutton, F.D., 399, 417 Swaminathan, S., 311 Tanaka, A., 249, 259, 746, 765 Tankersley, C.G., 272, 278, 287–288, 290
Author Index Tatsumi, K., 372, 379–380 Taylor B.J., 761, 776 J.H., 450, 499, 508 M.B., 531, 564 Temp, J.A., 674, 680 Tenney, S.M., 271, 288 Thach, B.T., 324, 349 Thomas A.J., 273, 288 P.S., 389, 392, 409, 415 Tian, Y., 459, 503 Tishler, P.V., 400, 418 Tonkovic-Capin, M., 582, 632 Trachsel, D., 536, 565 Tucker, D.H., 392, 417 Tuman, K.J., 483, 494 Tusiewicz, K., 703, 729 Tyler, M.K., 556, 570 Vangerven, M., 532, 565 Van Hemelrijck, J., 532, 565 Vargo, J.J., 536 Verbraecken, J., 398–399, 417 Vincent, H.H., 754, 771 Violet, J.M., 594, 643, 790 Vizek, M., 657, 678 Von Euler, C., 611, 646, 684 Wadstrom, C., 391, 415 Waizman, J., 390, 415 Walder, B., 450, 499 Waldhorn, R.E., 410, 422 Waldrop, T.G., 160, 203, 206–207
821 Wang W., 24–25, 57 X.G., 31, 63–64 Ward, D.S., 105, 130, 537, 566 Warley, A., 113, 128 Warner, D.O., 702, 704–705, 720, 722, 724, 726, 729, 734 Warren, P.M., 450, 499, 508 Way, W.L., 546, 568 Weil, J.V., 372–374, 379–381, 544, 566, 657, 678 Weiner, P., 390, 415 Weiskopf, R.B., 668, 683 Weitzner, S.W., 517, 560 Westphalen, R.I., 585, 638 White D.P., 325, 350, 373–374, 381 P.F., 449, 499, 520, 525–526, 561–562 Whitwam, J.G., 531, 564 Winnie, A.P., 553, 569 Wise, R.A., 398, 417 Wyon, N., 783–784 Yamamoto, M., 804, 812 Yamashiro, S., 106, 125 Yeager, M.P., 483, 491, 493 Yost, C.S., 589, 640 Young, T., 314, 338–339, 341, 399, 414 Zerah, F., 389, 416 Zhou, H.H., 545, 567 Zsigmond, E.K., 453, 500, 555–556, 570 Zuperku, E.J., 599, 626 Zwillich, C.W., 399, 417
SUBJECT INDEX
Abdomen effect of anesthesia on, 688–689 muscles of, effect of anesthesia, 689 surgical procedure, recovery from anesthesia, 752–753 Acetazolamide, 26, 28–31, 34, 38, 42, 365, 368, 372–373, 601 Acetylcholine, 4, 9–13, 41, 44, 46, 235, 241, 527, 572, 591, 593–594, 596, 779, 781–783 receptor, 9, 11, 572, 593–594, 596, 779, 783 Acidosis, putative chemoreceptor areas, 28 Acromegaly, obstructive sleep apnea, 316 Acute mountain sickness, 372–374. See also Mountain sickness acetazolamide, 372–373 almitrine bismesylate, 373 ginkgo biloba, extract from, 373 hypercapnic ventilatory response, 372 L-tryptophan, 374 progestational steroids, 374 sedative-hypnotics, 373–374 temazepam, 373–374 theophylline, 373 zolpidem, 373–374 Adenosine, 15, 161–162, 193–194, 364, 473, 608, 657, 802–804 Adenosine triphosphate, 4, 9, 11–15, 40, 46, 666, 781–782, 805 Adrenaline, 4, 36, 76, 198, 434, 475, 607, 626, 629, 673, 747, 795–796, 798 Advanced cardiac life support, 452
Afferent innervation lower airway/receptors, 234–238 upper airway/receptors, 226–229 Agonist/antagonist compounds, 473–474 Air hunger, 80 Airway, 225–259 afferent innervation, receptors lower, 234–238 upper, 226–229 cardiovascular responses, 249–250 central nervous system, 226 C-fiber endings, 236–237 clinical problems, 244–250 cough, 246–248 gas concentration of drug, 106–108 laryngospasm, 248 lower chemical stimulation, 240–244 integrative aspects of airway reflexes, 238–244 lung inflation, 238–240 mechanical stimulation, 240–244 slowly adapting receptor activation, 238–240 neuroepithelial bodies, 237–238 obesity, resistance with, 389 lower airway, 389–391 neck circumference increased, 389 upper airway, 389 obstruction of, 248–249 patency, maintenance of, 233–234 pressure alterations, 120–122 flow limitation, 122
823
824 Airway (continued ) genioglossal EMG, 122 negative airway pressure, 122 resistance, 122 protective functions, 229–233 impaired, in recovery from anesthesia, 748–749 pulmonary aspiration, 244–246 rapidly adapting receptors, 236 slowly adapting receptors, 235–236 upper, reflex responses from, 229–234 airway, defense, protection, 229–233 apnea, 231 larynx, 230–231 maintenance of airway patency, 233–234 nose, 229 pharynx, 232 Akaike’s information-theoretic criterion, pharmacodynamic interaction modeling, 137–138, 142–143, 151 Alfentanil, 477, 554 sevoflurane, interaction, 143–148 steady-state technique, 554 Almitrine bismesylate, 103, 124, 373–375, 658 Alpha-adrenergic agonists, antagonists, 476–481, 798–800 Alpha-amino-3-hydroxy-5-methylisoxazole4-propionate, 572, 591–593, 596, 605, 613–615, 617, 621, 623 Altitude, pulmonary edema. See High-altitude pulmonary edema American Society of Anesthesiologists, 516, 525–526, 532, 557 2-amino-5phosphonovalerat, 572, 605–606, 615 Aminophylline, 162, 755, 803–804 Amrinone, 755 Amygdala, opioids and, 442 Analgesia, 107–108, 428–440, 442, 445–490, 513–516, 525, 532, 540, 544, 546, 553, 555, 557, 559, 576, 741, 754, 757, 760–761, 802. See also Pain; specific analgesic agent central, 454 epidural, 425, 431, 436–437, 459–465, 474, 483–486, 488, 490, 754, 761 patient-controlled, 107, 383, 447–450, 453–454, 461–462, 473, 482, 486–487, 490 epidural, 436, 462, 490
Subject Index peripheral, 454 preservation of, 453 Anesthesia, 229–234, 245–250, 325–328, 425–652, 687–782. See also specific anesthetic agent airway protective functions, impaired, after, 748–749 atelectasis, postoperative, 754–755 brachial plexus block, 454, 457–458, 475, 479 cardiovascular function, effects of pain on, 437–438 central effects of, 571–652 chemical control of breathing after anesthesia, 750–752 chest wall mechanics, effects on, 694–709 chronic pain, 438 clinical significance, 755–758 control of breathing, postanesthetic residual drug effects on, 750–751 general, 687–735 hypoxemia late, treatment strategy for, 763–764 postoperative, late, treatment strategy for, 763–764 during sleep, postoperative, mechanisms of, 762–763 intrathecal, 444, 453–455, 457, 462, 467–473, 480, 482–483, 487–488 laryngeal airway maintenance function, impaired, 745–748 laryngeal edema, after extubation, 745 laryngospasm, during emergence from anesthesia, 745 lung function effects on, 714–721 impairment, recovery of, postoperative, 752–755 lung volume changes, postoperative, 752–754 nocturnal hypoxemia, late postoperative, 755–764 opioids, 440–454 control of breathing, 751 postoperative sleep disturbance and, 760 pain, effects of, 438–454 pain management, 438, 751–752 ventilatory effect, effects of pain, 438–454 pharyngeal obstruction analgesia with opioids, 741
Subject Index Anesthesia (continued ) body position, effects of, 744 head position, effects of, 744 mechanisms of, 739 neck position, effects of, 744 during recovery from anesthesia, 738–745 residual muscle paralysis, 742 surgical, anesthetic interventions, effects of, 742 pneumonia, postoperative, 754–755 postoperative respiratory dysfunction, 426–433 pulmonary function, effects of pain on, 436–437 recovery from, 737–777 regional, 425–512, 760 respiratory ciliary dysfunction, postoperative, 754 respiratory pump, normal function, 688–693 segmental high thoracic epidural, 463 sleep architecture, postoperative changes in, 758–762 sleep disturbance, postoperative, surgical stress and, 761 stress response, effects of pain on, 433–436 stridor, after extubation, 745 surgical trauma, effects of, 433–438 upper airway function, recovery from anesthesia, 738–749 upper airway mechanics, effects on, 709–714 Anesthetics, 233–237, 334, 460–468, 572–677, 695, 697, 706, 709–714, 721–722, 738–748. See also specific anesthetic agent fast synaptic neurotransmission, 583–596 gamma-aminobutyric acid, 594, 622 glutamate receptors, 591 glutamatergic excitation, block of, 622 glycine receptors, 595 halogenated volatile, 658 hypoglossal motoneuron depression, 626–629 excitability, 626 inhalational, 122, 573, 608–609, 628, 653–655, 659, 663–671, 676–677, 742, 750, 754 (See also specific inhalational anesthetic)
825 intramuscular, 447–448, 520, 527–528, 537, 753, 760 (See also specific intramuscular anesthetic) ligand-gated ion channels, 591–596 local, 246, 427, 435, 454–455, 460–468, 479, 481, 484–485, 488–489 (See also specific local anesthetic) membrane potential of respiratory neurons, 622–626 neuronal nicotinic acetylcholine receptors, 593 oscillator model, for central anesthetic effects on respiratory rate, 608 potassium channels, 589 anesthetic-induced, 626–629 respiratory minute ventilation, 574–580 respiratory neurons, 581–583 respiratory neurotransmission, 613–629 tandem pore acid-sensitive potassium channel (TASK), 6, 8, 13, 46–47, 52, 73–75, 86, 89, 262, 296, 514, 590–591, 627, 664, 673 volatile carotid body mediated ventilatory response, 658 effects of, 574, 615 minimum alveolar concentration of, 655 at subanesthetic concentrations, 658 voltage-gated ion channels, 584–591 voltage-gated potassium channels, anesthetic insensitivity, 589–590 voltage-gated sodium channels, 588 Angina, 801 Angiography, 553 Angiotensin converting enzyme, 247, 265, 280, 801 Angiotensin-converting enzyme, 247 gene, 265, 802 Angiotensin II, 800–801 Animal models, ventilatory behavior inheritance, 261–291 Antihypertensive agents, 476 Antitussive actions of opioids, 442–443 Anxiety, postoperative, 435 Anxiety disorders, breathing patterns with, 77 Anxiolysis, 477, 514–515 Aortic chemoreceptors, 159, 185–186 Apnea-hypopnea index, 263, 397–398, 408 obstructive sleep apnea (See Obstructive sleep apnea)
826 Apneusis, induction of, 579–580 Apolipoprotein E, 263–264, 280 Arginine vasopressin, 800–801 Arousal responses, 673–674 Arrhythmias, postoperative, 435 Artificial brainstem perfusion, 50, 115, 580, 657, 663, 669 Aspiration pneumonia, 748 Asthma cough-variant, 247–248 treatment, 443 Atelectasis anesthetic effects, 716–720 postoperative, 435 in recovery from anesthesia, 754–755 Atrial natruretic peptide, 400 Automatic control of breathing, 72 Autonomic nervous system dysfunction, in hypoventilation syndrome, 304 Autonomopathy, congenital central hypoventilation syndrome as, 295–312 Awake behavior, species differences in, 695 Background potassium channel, 44, 64–65, 683 Barbiturate, 513, 527–532, 579–580, 584, 587, 591–594, 596, 608, 710 Behavioral noise, 82–83 Behavioral responses, inhalational anesthetics, carotid body mediated ventilatory response, 671–674 Benzodiazepine, 104, 334, 373–374, 445, 513, 516–527, 553–554, 556, 607, 622–624, 747–748 opioids combined with, 553–554 Bicarbonate compensation, 367 Bier block, 455–456, 479 regional, intravenous, 455–456 Bispectral index, 161, 167, 181, 373, 475, 532, 535 Blood-brain barrier, 796, 798, 803 Blood pressure surges, in obstructive sleep apnea, 317 Blood vessels, putative chemoreceptor cell association with, 24–25 Body mass index in obesity, 384–385, 387–395, 397–398, 400, 410 Bootstrap analysis, 140–151 propofol, remifentanil, 148–151 sevoflurane, alfentanil, 143–148
Subject Index Bo¨tzinger complex, 598 Brachial plexus blocks, 457–458 axillary approach, 458 buprenorphine, addition of, 458 dyspnea, 457 forced vital capacity, 457 hemidiaphragmatic paresis, 457 hoarseness, 458 interscalene approach, 457 periclavicular approach, 458 regional analgesia, 457–458 Bradycardia, 478 Brain blood flow, 657–658, 676 Brainstem central chemosensitive areas, 53–55 evolutionary, 53 rostral ventral lateral medulla, 53–55 ventral medullary raphe region, 55 Brainstem respiratory network, 597–612 anesthetic-induced changes in rhythm, 608–612 gamma-aminobutyric acid, 606 glutamate, 603 glycine, 606 mammalian model, 597 neurotransmitters, in rhythm, pattern generation, 603–608 oscillator model, 608 pulmonary afferents, rate changes, 609 rhythm generation in adult mammal, functional model for, 601 tonic respiratory drive, 599 Breathing route, switching of, 233–234 Breuer-Hering inflation reflex, 238–239 Bronchoconstriction, 231 Bronchoscopy, 513 Bronchospasm, postoperative, 428 Bronchus, stimulation to, 240 Brown Norway rats, heritability studies, 273, 276, 278 Bulbospinal neuron, 164, 598, 601, 607, 614 Bupivacaine, 475, 753–754 Buprenorphine, 458, 473–474 Butorphanol, 473 Calcium-activated potassium channel, 29, 45, 589 Calcium channel, 23, 584, 586, 588, 596, 801 Calcium channel blockers, 801–804 Cancers, in obesity, 385 Carbon dioxide, rebreathing, 708–709
Subject Index Carbonic anhydrase, 26, 28–31, 43, 364, 372, 601 Carbonic anhydrase immunohistochemistry, 30 Cardiac surgery, 487–488 anticoagulation, 488 atelectasis, 487 forced vital capacity, 487 intrathecal morphine, 487–488 morbidity, pulmonary, 487 oxygenation, 487 peak expiratory flow rate, 487 persistent hypoxemia, 487 pneumonia, 487 respiratory failure, 487 sternotomy, 487 thoracic epidural analgesia, 488 thoracotomy, 487 Cardiovascular disease, in obstructive sleep apnea, 316–317 Cardiovascular drugs, 793–813. See also specific drug alpha-adrenergic agonists, antagonists, 798–800 calcium channel blockers, 801–804 catecholamine agonists, antagonists, 794–800 dopaminergic agonists, antagonists, 796–798 purinoceptor agonists, antagonists, 802–804 renin-angiotensin system, 800–801 Cardiovascular function, effects of pain on, 437–438 Cardiovascular responses to apnea, 249–250 Carotid body, 3–16, 29, 31, 38–39, 44, 49, 51, 75, 120, 158, 161, 167–171, 176–182, 267, 277, 301, 329, 358–359, 364–369, 372, 442, 444, 446, 573, 590, 593, 599–600, 653–669, 671, 674–677, 742, 750–751, 780–785, 795–800, 802, 804–805 cellular organization, 4–6 chemoreceptors, 593–594 denervation, 48–49, 51, 185–186 function, pathways altering, 181 glucose sensing in, 12 innervation, 4–6 neuromodulation in, 12–15 oxygen sensing, 781
827 type-I cells, 4–15, 29, 39, 653, 796–797 type-II cells, 4, 6, 14–15 ventilatory response mediation, inhalation anesthetics, 653–686 Carotid sinus nerve, 3–4, 9, 48, 104, 161, 170, 180–181, 186, 362–366, 594, 657, 668–669, 781, 796 Catecholamines, 4, 249, 362, 437, 761, 794–800 Central chemoreceptors, 21–69 anatomical location, 24–38 electrophysiological studies in vitro, 27 blood vessels, putative chemoreceptor cell association with, 24–25 brainstem central chemosensitive areas, 53–55 evolutionary, 53 rostral ventral lateral medulla, 53–55 ventral medullary raphe region, 55 breathing, 47–55 carbonic anhydrase immunohistochemistry, 30 c-fos immunohistochemistry, 32 in caudal mesencephalon, 33 in caudal ventrolateral medulla, 32 dissociated from neuronal firing, 34 in dorsal medulla, 32 hypercapnia, 34 negative regulator of expression, 33–34 not limited to specific cells, 34 at pontine level, 33 in rostral brain areas, 33 in rostral regions of fastigial nucleus, 33 in rostral ventral lateral medulla, 32 in rostral ventral medial medulla, 32 single-cell resolution, technique with, 36 technique variations, 34 time pattern of, 33 cholinergic neurotransmission, 40–41 CO2 microdialysis, anatomical location, 30 electrophysiological studies in vitro, 27 acetazolamide injection, 28 acidosis, putative chemoreceptor areas, 28 focal acidosis, mechanism causing, 28 phrenic activity, time courses of changes in, 29
828 Central chemoreceptors (continued ) rapidly permeating carbonic anhydrase inhibitors, 29 extracellular pH CO2, 38–42 intracellular pH, 38–42 regulation of, 43–44 ventilation, relation between, 40 functional location, 24–38 glia, extracellular pH, 43–44 glial-glial coupling, functional role of, 44 intracellular, role in central chemoreception, 38–39 intracellular pH CO2, 38–42 extracellular pH, membrane channels, 44–46 gap junctions, 42 relation between, 41–42 inward rectifying potassium channels, 45–46 locus coeruleus, 27 mechanism of, 38–47 medullary serotonergic raphe neurons, large arteries, association, 24–25 metabolic acid-base disturbances, 50–51 neuron excitability, role in control, 30–31 neuron-to-glial coupling, functional role of, 44 neurotransmitters, synthesis release, uptake, 44 nucleus ambiguous, 27 nucleus of solitary tract, 27 ongoing synaptic transmission, modulation of, 40–41 peripheral CO2 measurement, 24–26 pre-Bo¨tzinger complex, 27 raphe nuclei, 27 retrotrapezoid nucleus, 27 rostral ventral lateral medulla, 27 rostral ventral medial medulla, 27 tandem pore acid-sensitive potassium channel (TASK), 6, 8, 13, 45–47, 52, 73–75, 86, 89, 144–145, 262, 296, 514, 590–591, 627, 664, 673 tonic drive to breath, 51 congenital central hypoventilation syndrome, 52 rostral ventral lateral medulla, influence of, 52
Subject Index rostrolateral medulla, unilateral focal lesions in, 52–53 ventilation, extracellular pH, relation between, 40 ventilatory response to CO2, 47–50 dynamic end-tidal forcing, 50 hyperoxia to separate peripheral, central inputs, 47 modified Read rebreathing technique, 49 recovery of CO2 response after, 49 ventilatory/phrenic nerve response to CO2, abolition of, 48 ventral medullary surface areas, cooling of, 48 ventriculocisternal perfusion, 47 ventral medulla, intracellular acidosis, 38 Central nervous system, 226 in congenital central hypoventilation syndrome, 300–301 functional magnetic resonance imaging, 301 depressants, 586, 596, 608 obstructive sleep apnea and, 334–335 Cephalad diaphragm displacement anesthetic effects on, 699 diaphragm insertions, 699–700 tonic diaphragm activity, anesthesia effects on, 699 Cervical contusion injuries, respiratory plasticity and, 196–197 Cervical hemisection, respiratory plasticity and contralateral effects, 196 supraspinal effects, 196 Cervical strap, obstructive sleep apnea and, 332 C-fiber endings, 234, 236–237, 243–244 Chemical control of breathing after anesthesia, 750–752 Chemoreceptors, 20–55, 72, 74, 76, 82, 159, 173, 181, 185–186, 195–196, 268, 273, 327–329, 358–376, 401, 471, 544, 573, 575, 593–594, 600, 628, 671–677, 780–785, 795–796, 802–805. See also specific chemoreceptor aortic, 159, 185–186 chest wall, 87, 89, 107, 189, 233, 262, 267–269, 329, 383, 388, 404, 430–431, 457, 462, 468, 687–701, 706–712, 716, 718–722
Subject Index Chemoreceptors (continued ) clusters, 4–6 peripheral, 3–19 (See also Peripheral chemoreceptors) Chest wall chemoreceptors, 87, 89, 107, 189, 233, 262, 267–269, 329, 383, 388, 404, 430–431, 457, 462, 468, 687–701, 706–712, 716, 718–722 function, effect of anesthesia on, 720–721 loaded breathing, 707–709 motion, anesthetic effects on, 701–709 quiet breathing, 706–707 respiratory muscle activation, 701 diaphragm, 704–706 Cheyne-Stokes breathing, 86, 273, 797, 804 Children, breathing pattern of, 577 Chloral hydrate, 538–540 electromyelogram, 540 hypoventilation, 539–540 infants, sedation for, 538–539 upper airway obstruction, 540 Chloroform, 611 Chlorpromazine, 555 Cholecystectomy, anesthesia recovery, 752–753 Cholecystokinin, 608 Choral hydrate, 538–540 Chronic mountain sickness, 374–376. See also Mountain sickness Chronic obstructive pulmonary disease, 74, 80, 88, 172, 271, 282, 392, 405–407, 463, 517, 552 Ciliary dysfunction, postoperative, 754 Circumventricular organs, 800–801 Cleveland Family Study, 263–264 Clonidine, 452, 458, 477–480, 483, 799 addition of, 458 epidural, 478–479 Cold (flow) receptors, 227 Colonoscopy, 558 Compensatory neuromuscular mechanisms, improvement of, 333–334 Congenital central hypoventilation syndrome, 85, 295–312 afferent respiratory neural input, defective integration of, 303 animal models, 300 autonomic nervous system dysfunction, 304 chemosensory, metabolic inputs, central integration of, 303
829 clinical presentation of, 296 defined, 296–298 diagnosis, 296–298 one of exclusion, 297 pathophysiology, 298–299 genetic hypothesis, 299 mutations in, screening for, 299 physiologic abnormalities, 301–304 respiratory physiology laboratory, evaluation in, 297–298 structural central nervous system abnormalities, 300–301 functional magnetic resonance imaging, 301 ventilatory dysfunction, severity of, 296 voluntary breathing, deficit in, 301–302 Consciousness, level of, in recovery from anesthesia, 742 Conscious sedation, 513–516. See also specific sedative defined, 513 Continuous positive airway pressure, 333, 335, 739, 755, 764 in obesity, 406, 409–411 in obstructive sleep apnea, 333, 335 resetting of device, 404 Control of breathing, 109–110 postanesthetic residual drug effects on, 750–751 Coronary artery thrombosis, postoperative, 435 Cough, 246–248 angiotensin-converting enzyme inhibitor, 247 chronic cough, 246–248 cough-variant asthma, 247–248 differential diagnosis, 246 gastroesophageal reflux, 247 index of disease, 246 post-nasal drip syndrome, 248 slowly adapting receptors, role of, 238–240 C-reactive protein, 385, 434 Crossed phrenic motor output, spinal cord injury, respiratory plasticity and, strengthening, 193–195 Cystoscopy, 513 Daytime hypercapnia, with obesity, prevalence of, 402 Decerebrate, 580–583, 608, 613, 617, 622, 625, 668, 710
830 Defensive reflexes airway, 246 larynx, 231 Dejours phenomenon, 273 in inheritance of ventilatory traits, 278 Delayed expiratory emptying, 578–579 Delirium, postoperative, 757–758 Delta receptor, 440, 475 Demerol, 555 Denervation, sensory, respiratory plasticity and, 185–186 Depression, postoperative, 435 Desflurane, 574–575, 584, 655, 660, 667–668 Developmental hyperoxia, carotid denervation, distinguished, 186 Developmental respiratory plasticity, 174–187 defined, 174 hypercapnia, 182–184 hyperoxia, 179 intermittent neonatal hypoxia, 178–179 metabolism, 184–185 neonatal maternal separation, 186–187 prenatal hypoxia, 175–176 sensory denervation, 185–186 sustained neonatal hypoxia, 176–178 Dexmedetomidine, 477–478, 799 Dextromethorphan, 481 Dezocine, 473 Diabetes mellitus, in obesity, 385 Diaphragm anesthetic effects on, 688–689, 704–706 Diaphragmatic descent, effect of anesthesia on, 690 Diazepam, 122, 514, 516–518, 520–521, 524–527, 553–554, 556, 624, 710 effect on ventilatory drive, 518–518 electromyelogram, 517 intramuscular, 520 intravenous, 520, 624 oral, 516–517, 520 parenteral, 517–520 rebreathing technique, 517 Digitalis, 805 2, 3-dihydroxy-6-nitro-7-sulfamoylbenzo(f)quinoxaline, 572, 605–606, 615 Dilator muscle activation in obesity, 401 in obstructive sleep apnea, 327 Diltiazem, 802 Dimorphism, sexual, inheritance of ventilatory traits, 282
Subject Index Diphenhydramine, 538–539, 556 alfentanil, 538 moderate sedation, 538 opioid, combined with, 538 opioid-related side effects, treatment of, 538 Dipyridamole, 802–804 Discomfort, respiratory, breathing and, 79–82 air hunger, 80 breathing effort or work, 80 chest tightness, 80 dyspnea, 79–82 afferent sources of perception, 82 cerebral correlates, 82 emotional disturbance, elicitation of, 80 inappropriate perceptions of, 81–82 influence on volitional drives to breathe, 80–81 intensity of, unpleasantness of, relationship between, 80 unpleasantness of, intensity of, relationship between, 80 voluntarily activation of breathing muscles, circumstances of, 81 Disinhibition, tonic excitatory drive, 85 Diurnal hypercapnia, with obesity, 406–407 Dizocilpine, 605, 624 Domperidone, 372, 797–798 Dopamine, 4, 9, 12, 36, 43, 160–162, 168, 176, 180–181, 198, 209–210, 216, 226, 251, 276, 280, 304, 366–367, 369, 372, 380, 537, 656–658, 781–782, 795–798, 802, 805 presynaptic effect of, 12–13 Dopamine agonist, 796–798 Dopamine antagonist, 162, 798 Dopamine receptor, 9, 364, 797 Dorsal neural network, 79 Dorsal respiratory group, 443, 582 Dorsal rhizotomy, respiratory plasticity and, 197–198 acute, 197 bilateral thoracic, 198 chronic, 197–198 Dose-dependently depressed, 581–582 Doxapram, 103, 366, 373, 553, 755 meperidine, compared with, 553 D2 receptor, 12, 364, 366–367, 798 Drive receptors, 227 Droperidol, 537–538, 555, 589–590, 751 endogenous dopamine, blockade of, 537
Subject Index Droperidol (continued ) with fentanyl, 555 hypoxic ventilatory response, 537 opioid, combined with, 537 D-tubocurarine, 781, 786 Dual-isohypercapnic technique, 117, 558 Dynamic analgesia, 489 Dynamic end-tidal forcing system test, 118 in ventilatory response to CO2, 50 Dynamic hypercapnic response pharmacodynamics model, 120 Dysphoria, 473 Dyspnea, 79–82 afferent sources of perception, 82 cerebral correlates, 82 EC50, 108, 520–521, 550, 584 Edema laryngeal, after extubation, 745 pulmonary, 89, 117–118, 134, 262, 358–360, 376, 520, 528, 531, 544, 603, 622, 688, 690, 692, 698–701, 716–720, 722, 801 (See also High-altitude pulmonary edema) Elastic behavior, lung, anesthetic effects on, 720–721 Electrolyte imbalance, postoperative, 435 Electromyelogram, 390, 397, 709 Emotional influences on breathing, 75–84 perception of music, 82–84 behavioral noise, source of, 82–83 as ‘‘mood enhancer,’’ 83–84 respiratory cycles, effects of musical stimulation, 83 perception of respiratory discomfort, 79–82 afferent sources of perception, 82 air hunger, 80 breathing effort or work, 80 cerebral correlates, 82 chest tightness, 80 dyspnea, 79–82 emotional disturbance, elicitation of, 80 inappropriate perceptions of, 81–82 influence on volitional drives to breathe, 80–81 intensity of, unpleasantness of, relationship between, 80
831 unpleasantness of, intensity of, relationship between, 80 voluntarily activation of breathing muscles, circumstances of, 81 psychological influences, 76–79 dimension model, 77–78 dorsal neural network, 79 emotion perception, cerebral correlates of, 78–79 expression of, effect on changes in breathing, 76–77 functional role, 78 self-regulation, involvement in, 78 specific breathing behaviors, distinguishment of psychological derangements, 77–78 specificity model, 77–78 ventral neural network, 79 Emotion perception, cerebral correlates of, 78–79 Endocrinopathy, 316 obstructive sleep apnea, 316 Endoscopy, 513, 553 Endothelin-converting enzyme-1, 276 Endotracheal tube, 443, 741, 744–746, 748, 754 Enflurane, 241, 528, 574–575, 581–582, 584–585, 588, 591–593, 595, 611, 621, 654–655, 658–663, 668–669, 704, 710, 742–743, 752, 754 Environmental factors, in obesity, 385 Epidural analgesic, 425, 431, 436–437, 459–465, 474, 483–486, 488, 490, 754, 761. See also Epidural blockade; Epidural opioids; specific analgesic Epidural blockade with local anesthetics, 461 block density, 461 block level, 461 hypotension, 462 local anesthetics, 462–465 motor weakness, 462 Epidural opioids, 465 biphasic respiratory depression, 465 bupivacaine, 466–467 fentanyl, 466 lipophilic opioids, 466 morphine, 465–466 neuraxial opioids, 465 sufentanil, 466–467 thoracotomy, 467
832 Epinephrine, 4, 36, 76, 198, 434, 475, 607, 626, 629, 673, 747, 795–796, 798 Epistasis, inheritance of ventilatory traits, 280–282 Erythrocytosis, 375 Ether, 236, 590, 611, 706 Etomidate, 584, 592, 594, 596 Eucapnic obesity, with obesity, 393 Excitatory postsynaptic potentials, 604, 625 Exercise, 122 hyperapnea, 89–90 Expiration reflex, 231 Expiratory muscles, phasic activity of, anesthesia effects on, 700–701 Expiratory neuron, 86, 581, 600, 613, 616, 618–622, 624, 785 inspiratory neuron, distinguished, 618–620 Expiratory reserve volume, in obesity, 388 Expression of emotion, effect on changes in breathing, 76–77 Extracellular pH CO2, central chemoreceptors, 38–42 intracellular pH, 38–42 Failure to thrive, 435 Fast-acting neurotransmitters, 8–11 Fast synaptic neurotransmission, anesthetic effect, 583–596 Fat distribution, in obesity, 385 Fenoldopam, 798 Fentanyl, 109, 443–453, 465–467, 469, 471–474, 490, 548–550, 552, 554–555, 557, 751, 754, 761, 802 droperidol combined with, 555 effect on ventilatory drive, 549 high lipid solubility, 548 isohypercapnic technique, 548–550 midazolam, interaction between, 554 morphine, compared with, 548 premedicating with, 557 rebreathing technique, 548–550 Field block, 456. See also local anesthesia Flumazenil, 104, 526–527 Flurazepam, 622–624 Focal acidosis, mechanism causing, 28 Forced vital capacity, postoperative, 431 Functional magnetic resonance imaging, 52, 75, 82, 86 Functional residual capacity, 695–701, 720 cephalad diaphragm displacement, 699 diaphragm insertions, 699–700
Subject Index expiratory muscles, phasic activity of, 700–701 tonic diaphragm activity, 699 inward rib cage displacement, 697 expiratory muscles, phasic activity of, 699 thoracic curvature, 698 tonic inspiratory muscle activity, 697–698 oxygenation impairment due to reduction of, 763 postoperative, 431, 752–753 Galanin, 608 Gall bladder disease, in obesity, 385 Gamma-aminobutyric acid, 13, 367, 579, 584–585, 594–596, 601–604, 606– 608, 613–626, 630, 657–658, 663 receptor, 579–580, 594, 606–607, 625, 630 Gas exchange, lung, anesthetic effects on, 716–720 Gastric tube placement, 743 Gastroesophageal reflux, cough and, 247 Gene effects, ventilatory traits, in rodent models, 276–279 Gene loci, identification of, 271 General anesthesia, 436, 514–515, 687–735. See also Anesthesia; see also specific anesthetic agent central effects of, 571–652 chest wall, 688–692 components, 688 mechanics, 694–709 normal chest wall mechanics, 690 lung mechanics, effects on, 714–721 respiratory pump, normal function, 688–693 upper airway, 692–694 larynx, 693–694 mechanics, effects on, 709–714 muscle activation, 710–712 normal upper airway, 693 palate, 692 pharynx, 692 Genetic inheritance of ventilatory traits, 261–291 animal models, 261–266 physiogenetic map, ventilatory behavior, 279–282 in rodent models, 271–273, 276–279 in small animals, 266–271 strength of inheritance, 274–276
Subject Index Genioglossus, in obstructive sleep apnea, 329–332 Ginkgo biloba extract, in mountain sickness, 373 Gkytanate, 367 Glia, extracellular pH, 43–44 Glucose, 4, 12–13, 15, 44, 316–317, 385, 461, 653 Glutamate, 591, 603, 626–627, 658 Glutamatergic, 622 Glycine, 473, 592, 603, 606, 613, 626 receptor, 595–596, 601, 603, 606–607, 663 Guidelines for Sedation and Analgesia by Non-Anesthesiologists, 516, 525–526, 532, 557 Halogenated volatile anesthetics, 658. See also volatile anesthetics carotid body mediated ventilatory response, 658 Halothane, 43, 45, 531, 535, 574–595, 608, 610–612, 617–630, 654, 658–677, 696, 698, 700–711, 717–718, 720, 742, 745, 750, 754 Heart disease, in obesity, 385 Heart failure, 375 Heme protein, 8 Hemisection, spinal, respiratory plasticity, 189–192 breathing at rest, 189–190 contralateral effects, 196 crossed phrenic activity, 191–192 crossed phrenic pathways, 190–191 crossed phrenic phenomenon, 190–191 minute ventilation, 189 Hemothorax, postoperative, 428 Hering-Breuter reflex, 610–612 High altitude, 357–382 acute hypoxic ventilatory response, 361–363 acute mountain sickness, 372–374 chronic mountain sickness, 374–376 hypercapnic ventilatory response, increases with acclimatization, 372 hypoxic desensitization, 369 hypoxic ventilatory response, 363 time domains of, 360–371 intermittent hypoxia, 369–371 periodic breathing during sleep, 374 short-term depression, 363 short-term potentiation, 362-363 sustained hypoxia responses, 363–369
833 ventilatory acclimatization, 365 ventilatory deacclimatization, 368 High-altitude pulmonary edema acute, 372–374 chronic, 374–375 Hippocampal pyramidal neurons, 586–588 Human ether-a-gogo-related gene, 8 Hydromorphone, 448–449, 454, 467, 500 2-hydroxyethyl-piperazineethane sulfonic acid, 39 5-hydroxytryptophan, 4, 13, 43–45, 157, 161, 163–168, 170, 186, 190–195, 197–198, 226, 236–237, 277, 362, 367, 376, 473, 475, 572, 607–608, 626–629, 673 Hydroxyzine, 555 Hyperbola model, 118 Hypercapnia, 110–117, 182–184, 364, 368, 600, 667, 742, 750–752, 797, 800 with acclimatization, 372 in mountain sickness, 372 nocturnal, in obesity, 404 in obesity, 392–399, 409–410 permissive, 182–183 respiratory plasticity and, 172–174, 182–184 acute intermittent, 172 acute sustained, 172 chronic intermittent, 174 chronic sustained, 173 Hypercoagulable state, postoperative, 435 Hyperdynamic circulation, effect of pain, 434 Hyperglycemia, postoperative, 435 Hyperlipidemia, in obesity, 385 Hyperoxia, 171–172, 179, 674 Hyperpolarization, 599, 628 Hypersomnolence, 314–315, 408 Hypertension, 801 in obesity, 385 postoperative, 435 Hyperventilation-induced hyperapnea, 368 Hypocapnia, 365, 368, 580, 600, 803 Hypoglossal motoneuron depression, 626–629 excitability, 626 Hypotension, 478, 804 Hypothyroidism, 316 obstructive sleep apnea and, 316 Hypoventilation syndrome, central, congenital, 295–312
834 Hypoxemia, 558, 742, 797 nocturnal, 755–764 with obesity, 392–399 postoperative, 428, 763–764 during sleep, 762–763 (See also Nocturnal) treatment, 763–764 Hypoxia, 8, 167, 357–376, 655, 659–660, 664, 667, 673–674, 676–677, 797, 804. See also Hypoxic ventilatory response central nervous system depressant mechanisms, 168 defense mechanism, 653 intermittent, 163, 168, 369–371 with long-duration hypoxia, 170–171 with moderate-duration hypoxia, 170 neonatal, 178–179 with short-duration hypoxia, 169 postoperative, 435 respiratory plasticity and, 158–171 acute, 159 hypoxic ventilatory depression, 161–162 post-hypoxia frequency decline, 162–169 short-term depression, 162–169 short-term potentiation, 159–161 sustained, 167 ventilatory acclimatization to, 167–168 Hypoxia-driven ventilation, 661 Hypoxic chemosensitivity, neuromuscular blocking agents, interaction between, 783 Hypoxic chemotransmission, 11 Hypoxic desensitization, 369 Hypoxic pulmonary vasoconstrictor response, 376 Hypoxic ventilatory depression, 120, 655, 658–659, 663, 676 Hypoxic ventilatory response, 117–120, 358–360, 363, 370, 372, 375, 750–752, 798, 802 blunting of, 369 genetic control of, 278–279 sensitivity, 666–667 time domains of, 360–371 Immunoincompetence, postoperative, 435 Incentive spirometry, 755 Inductance plethysmography, 108
Subject Index Inhalational anesthetics, 122, 573, 608–609, 628, 653–655, 659, 663–671, 676–677, 742, 750, 754. See also specific inhalational anesthetic carotid body mediated ventilatory response, 653–686 Inheritance of ventilatory traits, 261–291 animal models, 261–266 physiogenetic map, ventilatory behavior, 279–282 in rodents, 271–273, 276–279 in small animals, 266–271 strength of inheritance, 274–276 Inhibitory ligand-gated ion-channels, 663–664 Inhibitory postsynaptic potential, 572, 595, 606–607, 622, 625 Inspiration, effect of anesthesia on, 693–694 Inspiratory bulbospinal neuron, 614 Inspiratory neurons, expiratory neurons, difference between, 618–620 Integration of airway reflexes, 238–244 Interaction models, pharmacodynamic, 134–137 mechanism-based approach, 135–136 modeling interaction, 136–137 model selection, 137–140 Richards model, 134–135 Intercostal nerve blocks, 458–460 cryoanalgesia, 459–460 functional residual capacity, 459 Joule-Thomson effect, 459 local anesthetics, 458–460 systemic opioid analgesia, compared with, 459 thoracotomy, 459 Internal intercostals, effect of anesthesia on, 701–702 Interpleural analgesia, 460–461 bupivacaine, 461 local anesthetics, 460–461 open cholecystectomy, 460 pneumothorax, 460 systemic toxicity, 460–461 thoracotomy, 460 unilateral bronchospasm, 461 Intra-articular analgesia, 456 Intracellular pH central chemoreceptors, 38–42 extracellular pH, membrane channels, 44–46 gap junctions, 42
Subject Index Intramuscular anesthetics, 527, 537. See also specific intramuscular anesthetic Intrathecal anesthesia alfentanil, 472 clonidine, 480 CSF, 469 dose-related analgesia, 469 expiratory function, 468 fentanyl, 469, 471–473 hydrophobicity, 472 hypoxia, ventilatory response, 469–471 inspiratory function, 468 intrathecal morphine sulphate, 472–473 lipid-soluble opioids, 471–472 with local anesthetics, 468 meningeal permeability, rank order for, 472 meperidine, 469 morphine, 469, 472–473 opioids, 469 oxyhemoglobin desaturations, 469 patient-controlled analgesia, morphine, 473 postoperative pain management, 468 respiratory depression, 469, 471–472 sufentanil, 469, 471–473 ventilatory mechanics, 468 Intrathoracic blood volume, effect of anesthesia on, 696–697 Inward rectifying potassium channel, 45–46, 55, 589–590, 679 Inward rib cage displacement, thoracic curvature, functional residual capacity, anesthesia effects on, 698 Ion channels, 40, 44, 46, 55, 181, 653, 663 Isocapnic conditions, maintenance of, 117–118 Isocapnic hypoxia, 359, 371, 675–676, 798 phrenic responses, 180 Isoflurane, 45, 118, 122–123, 230, 335, 453, 574–576, 581, 584–595, 628, 654–655, 658–669, 673–677, 696, 706, 712, 714–715, 745, 754, 759–760 Isoproterenol, 798–799 Joule-Thomson effect, 459 Kappa receptor, 442, 473–475 Ketamine, 536–537, 576–579, 584, 587, 589–593, 596, 696, 706, 710
835 benzodiazepine combined with, 556 opioid combined with, 556–557 propofol combined with, 557 ventilatory effects, 537 Ketanserin, 13, 163, 197 Knock-out approach, in inheritance of ventilatory traits, 276–278 Krebs cycle, 666 Laparoscopic surgery, anesthesia recovery, 752 Large conductance potassium channel, 6–8, 670 blockade of, 7–8 Laryngeal airway maintenance function, impaired, 745–748 Laryngeal edema, after extubation, 745 Laryngeal receptors, 227 Laryngospasm, 248 during emergence from anesthesia, 745 Larynx, 230–231 closure reflex, 231–232 cough reflex, 231–232 defense reflexes, 231 general anesthesia, effect on, 693–694 swallowing, 231 Lateral reticular nucleus, 32 Learned respiratory behaviors, 88–90 blood-gas disturbances, avoidance of, 90 exercise hyperapnea, 89–90 learned feedforward responses, 90 muscle activity, modulation of, 88–89 Left ventricular hypertrophy, 400 Leptin, 265, 276–277 Levels of sedation, definitions of, 515 Lidocaine, 234, 250, 455–456, 464, 745 Ligand-gated ion channel, 11, 583–584, 591–596, 622 Linear-in-saturation pharmacodynamics model, 118 Lipolysis, postoperative, 435 Loaded breathing, anesthetic effects on, 707–709 Local anesthetics, 246, 427, 435, 454–455, 460–468, 479, 481, 484–485, 488–489. See also specific local anesthetic cardiovascular depression, 455 epidural blockade with, 461 hypercarbia, 455 lumbar epidural lidocaine, 455 resting ventilation, 455
836 Local anesthetics (continued ) spinal, epidural, 455 systemic toxicity, 455 Locus coeruleus, 27, 33–34, 37, 42–46, 52, 205, 476, 601, 628 Long-duration hypoxia, chronic intermittent hypoxia with, 170–171 Lower airways chemical stimulation, 240–244 integrative aspects of airway reflexes, 238–244 lung inflation, 238–240 mechanical stimulation, 240–244 slowly adapting receptor activation, 238–240 L-tryptophan, in mountain sickness, 374 Lung abscess, 245 elastic, increases in, effect of anesthesia, 697 inflation of, 238–240 mechanics of, anesthetic effects on, 714–721 postoperative function, 752–755 total capacity of, 388–390, 464, 721 Lung volume, 72, 74, 78, 87, 235, 323, 325, 388–389, 402, 428, 430–431, 435–436, 443, 447, 462, 466, 487–488, 601, 688–691, 695, 697, 705, 710, 714, 718, 720–722, 752–754, 763 Magnetic resonance imaging, 42, 75, 78, 81–82, 86, 188, 297, 301, 316, 324, 693, 746, 755. See also Functional magnetic resonance imaging Maternal separation, neonatal, respiratory plasticity and, 186–187 immobilization with, 186 stress with, 186–187 Mechanical stimulation of airway, 240–244 Mechanism-based pharmacodynamic interaction models, 135–136 Melanocortin, 264–265, 385 Membrane hyperpolarization, 625 Membrane potential of respiratory neurons, anesthetic effect, 622–626 Meperidine, 445–449, 467, 469, 524–525, 546–548, 550, 552–556, 558, 721 effect on ventilatory drive, 547 endoscopic procedures, 546 isohypercapnic technique, 546
Subject Index moderate or deep sedation, 546 morphine, compared with, 546–548 normeperidine, 546 rebreathing technique, 546–548 Mesencephalic reticular formation, opioids and, 442 Metabolic alkalosis, 445 Metabolic syndrome, in obesity, 385 Metabolism, respiratory plasticity and, 184–185 Metabotropic glutamate receptor activation, 605–606 Metenkephalin, 608 Methadone, 448 Methohexital, 516, 528–529, 531–532, 589–590, 696, 706 hypoxemia, 528 intravenous, 528–529 moderate, deep sedation, 528 pediatric patients, 528 rectal, 528 spontaneous ventilation, effect of, 529 Methoxyflurane, 587, 611, 696 Midazolam, 445, 514, 521, 526–527, 556, 558, 589–590, 754 diazepam, compared, 521–525 effect on ventilatory drive, 522–523 fentanyl, interaction between, 554 intravenous, 521 isocapnic rebreathing technique, 521–524 obstructive sleep apnea and, 335 oral, 521 premedicating with, 557 Minimum alveolar concentration, 580, 583, 591, 616–622 Mitochondrial membranes, complexes I or II of respiratory chain in, 666 Moderate-duration hypoxia, chronic intermittent hypoxia with, 170 Modulation, defined, 156 Monge’s disease. See Hypoxic ventilatory response Monte Carlo simulation in pharmacodynamic interaction modeling, 139 Morphine sulfate, 109, 113, 117, 160–161, 191–192, 440–441, 443–451, 453–454, 456, 461, 465–479, 482, 486–488, 490, 516, 540–546, 548, 550, 552–553, 555–557, 741, 751–753, 760, 775, 802 age-related difference, 545 effect on ventilatory drive, 541–543
Subject Index Morphine sulfate (continued ) ethnicity, 545–546 gender, 545 hypoxic ventilatory response, 544 intravenous, 544 mu receptor, 540 opioids and, 440 postoperative, 760 rebreathing technique, 544 resting ventilation, 540 steady-state technique, 540–544 Motility in gut, pain and, 438 Mountain sickness, 372–376 acetazolamide, 372–373 acute, 372–374 almitrine bismesylate, 373 chronic, 374–376 ginkgo biloba, extract from, 373 hypercapnic ventilatory response, 372 L-tryptophan, 374 progestational steroids, 374 sedative-hypnotics, 373–374 temazepam, 373–374 theophylline, 373 zolpidem, 373–374 Mount Everest, 370. See also High altitude; Mountain sickness Mucous secretion, 231 Multi-modal analgesia, 489 Mu-receptor, 440–443, 474–475, 540 Muscimol, 41, 616, 619 Music perception behavioral noise, source of, 82–83 as ‘‘mood enhancer,’’ 83–84 respiratory cycles, effects of musical stimulation, 83 Myocardial infarction, postoperative, 435 Nalbuphine, 442, 453–454, 473–474 Naloxone, 104, 375, 440, 451–454, 472, 474–475, 489, 552, 577–578 antagonist of, 452 chronic obstructive pulmonary disease, 552 hypercarbia, 552 hypoxia, 552 hypoxic ventilatory drive, 552 opioids and, 440 prophylactic infusion, 489 reversal with, 474 Nasal continuous positive airway pressure, 764
837 in obesity, 404 in recovery from anesthesia, 739 Nasal receptors, 227 Negative nitrogen balance, postoperative, 435 Negative-pressure reflex, 249 Negative upper airway reflex, 233 Neonatal maternal separation, respiratory plasticity and, 186–187 immobilization with, 186 stress with, 186–187 Neostigmine, 479, 483, 744, 785 Neuraxial blockade, 435, 461–473 bupivacaine, 468–469 CSF, 467 delayed ambulation, 462 epidural analgesia, 462–465 epidural anesthesia, 462–464 expiratory reserve volume, 464 fentanyl, 467 functional residual capacity, 464 hydromorphone, 467 hypercapnia, 468 inspiratory capacity, 464 isocapnic hypoxia, 468 lidocaine, 464 lipid-soluble opioids, 467 meperidine, 467 morphine, 467 neuraxial opioids, 467 nonsteroidal anti-inflammatory drugs, supplementation with, 465 patient-controlled epidural analgesia, 462 peak expiratory flow rate, 464 pentazocine, 464 sameridine, 469 spinal anesthesia, 468 sufentanil, 467 thoracic epidural analgesia, 464 thoracic epidural anesthesia, 463 total lung capacity, 464 urinary retention, 462 vital capacity, 464 Neuroepithelial bodies, 237–238 Neurokinins, 607 Neuromuscular blocking agents, 779–813 carbon dioxide sensing, 780–781 blood-brain barrier, 780 carotid body, 780 central nervous system depressants, 780 hypercapnic ventilatory response, 780
838 Neuromuscular blocking agents (continued ) minute volume, 780 tidal volume, 780 diaphragm, 785–787 contraction force, 786 hypoxic chemosensitivity, interaction between, 783 oxygen sensing, 781–785 acetylcholine, 781–782 adenosine triphosphate, 781 atracurium, 782 blood-brain barrier, 781 carotid body, 781–787 cholinergic transmission, carotid body, interference with, 784 dopamine, 781 hypoxia, 783 hypoxic chemosensitivity, 783 hypoxic ventilatory response, 782–783 nitric oxide, 781 pancuronium, 782 substance P, 781 vecuronium, 782–783 regulation of breathing, 780–785 respiration, effects of curare on, 780 respiratory pump function, 785–789 upper airway, 785–789 Neuronal discharge pattern, 601–603, 607 Neuronal excitability, 365, 584, 586–587, 589, 608, 625, 627, 673 Neuronal nicotinic acetylcholine receptors, anesthetic effect, 593 Neuronal nitric oxidase synthase, 676 Neuropeptide Y, 264, 607 Neuroplasticity, respiratory, 155–223 cervical hemisection contralateral effects, 196 supraspinal effects, 196 crossed phrenic pathways, spontaneous motor recovery, 192–193 developmental plasticity, 174–187 hypercapnia, 182–184 hyperoxia, 179 intermittent neonatal hypoxia, 178–179 metabolism, 184–185 neonatal maternal separation, 186–187 prenatal hypoxia, 175–176 sensory denervation, 185–186 sustained neonatal hypoxia, 176–178 hypercapnia, 172–174
Subject Index acute intermittent hypercapnia, 172 acute sustained hypercapnia, 172 chronic intermittent hypercapnia, 174 chronic sustained hypercapnia, 173 hyperoxia, 171–172 hypoxia, 158–171 acute, 159 brief intermittent, 163 chronic intermittent, 168 hypoxic ventilatory depression, 161–162 post-hypoxia frequency decline, 162–169 short-term depression, 162–169 short-term potentiation, 159–161 sustained, 167 location of, 156–158 with long-duration hypoxia, 170–171 with moderate-duration hypoxia, 170 modulation, contrasted, 156 respiratory gases inducing, 158–174 sex hormones, 187 with short-duration hypoxia, 169 spinal cord injury, 187–198 cervical contusion injuries, 196–197 crossed phrenic pathways, 192–195 dorsal rhizotomy, 197–198 hemisection, 189–192 serotonin, 192 Neutral endopeptidase, 276 Nicotinamide adenine dinucleotide phosphate, 8, 277 Nimodipine, 802 Nitric oxide, 804–805 Nitric oxide synthase, enzymes, 265–266 Nitroprusside, 800, 804–805 Nitrous oxide, 453–454, 459, 513, 577, 592–594, 655, 674, 703–704 N-methyl-D-aspartate, 481–482, 579, 591–593, 605, 613–615, 621, 624 bupivacaine, intrathecal ketamine added to, 482 dextromethorphan, 481 dynamic analgesic, improvement of, 482 ketamine, 481 Nociceptive impulses, effect of pain, 433–434 Nocturnal hypercapnia, with obesity, 402 Nocturnal hypoxemia, postoperative, 755–764 oxygenation, 755–756 oxygen desaturation index, 756 defined, 755
Subject Index Nocturnal hypoxemia, (continued ) polysomnographic assessment, 756 wakefulness, 755 Nocturnal oximetry, with last postoperative hypoemia, 764 Non-invasive mechanical ventilation, in obesity, 410–412 Non-rapid eye movement sleep, postoperative, 761 Non-steroidal anti-inflammatory drugs, 450, 465, 482–483, 489 Norepinephrine, 4, 36, 176, 198, 434, 475, 607, 626–627, 629, 673, 795–796, 798 Normocapnia, 580 Normoxia, 358–359, 371 Nose, 229 depression of breathing, 230–231 irritation of, 229–230 stimulation of, 230–231 Nucleus ambiguous, 673–673 Obesity, 313–316, 323–325, 383–422 airways resistance in, 389 lower airway, 389–391 neck circumference increased, 389 upper airway, 389 anesthetic effects, 696 apnea, defined, 399 apnea-hypopnea index, 397–398, 408 bilevel positive airway pressure, 404, 406, 410–412 body mass index, 384–385, 387–395, 397–398, 400, 410 body surface area, 409 cancers, 385 chemosensitivity, therapy with, 404 chronic obstructive pulmonary disease, 405, 407 compliance, 388 continuous positive airway pressure, 406, 409–411 CO2 production, 391–392 daytime hypercapnia, prevalence of, 402 defined, 384 diabetes mellitus, 385 dilator muscles, activation of, 401 diminished inherent chemosensitivity, 405 diurnal hypercapnia, 406–407 DLCO, 392–393 electromyelogram, 390, 397
839 environmental factors, 385 etiology, 384–385 eucapnic obesity, 393 expiratory reserve volume, 388 familial chemosensitivity, role of, 405–406 fat distribution, 385 fatty tissue, loss of, 408 forced vital capacity, 388, 398–399 functional residual capacity, 388–389, 408 gall bladder disease, 385 gender effects in, 396–397 health effects, 385–388 heart disease, 385 hypercapnia, 392–399, 409–410 hyperlipidemia, 385 hypertension, 385 hypoxemia, 392–399 intermittent nocturnal hypercapnia, 404 lung volume, 402 massive, 407 maximum voluntary ventilation, 391, 398–399 metabolic syndrome, 385 nasal continuous positive airway pressure, resetting of device, 404 neuromuscular drive, increase in, 390–391 nocturnal hypercapnia, 402 non-invasive mechanical ventilation, 410–412 obesity hypoventilation syndrome, 391, 406 obstructive sleep apnea, 313–316, 323–325 with hypercapnia, 402–407 obstructive sleep apnea-hypopnea syndrome, 399–402 defined, 399 epidemiology, 384, 399 eucapnic obesity, 397 genetics of, 384–385 hypercapnic obesity, 398 nasal mechanical devices, 409–412 pathogenesis, 401 treatment, 408–412 weight loss, effect of, 408 O2 consumption, 391–392 overlap syndrome, 412 PaCO2, effect of weight loss on, 408–409 Pickwickian syndrome, 398
840 Obesity (continued ) polysomnography, 396 prevalence of, 384 respiratory muscle function, 390–391 serum cholesterol levels, 400 single breath diffusion capacity, 392–393 treatment, 408–412 ventilatory control disorders with, 399–407 ventilatory response, effect of weight loss on, 408–409 waist-hip ratio, 400 weight loss, improvement in lung mechanics with, 388–389 Obstructive sleep apnea, 248–249, 313–356, 741 acromegaly, 316 airflow, maximal inspiratory, 321–322 anatomic factors, 322–325 apnea-hypopnea index, 320–321 blood pressure surges, 317 cardiovascular disease, 316–317 central nervous system depressants, 334–335 cervical strap, 332 cervical structures, 322–321 clinical risk factors, 314–317 continuous positive airway pressure, 333, 335 critical pressure, 319–328 lowering of, 333 EMG activity of muscles, 328–330 endocrinopathy, 316 endotracheal extubation in, 743 epidemiology, 314–317 factors, modeling of, 318–320 familial factors in, 263–266, 315 gender and, 315 genioglossus decreases in activity, 329–330 role of, 330–332 with hypercapnia, obesity and, 402–407 hypothyroidism, 316 monitoring technology, 314 neuromuscular compensatory mechanisms, 325–327, 333–334 neuromuscular factors, 325–333 neuromuscular reflexes, 325 upper airway muscles, 330 with obesity, 313–316, 323–325, 399–407 oropharyngeal muscles, 333 pathogenesis, 317–333
Subject Index pathophysiology, 317–322 peri-operative setting, 334–335 pharynx, site of obstruction, 317–319 Pickwickian syndrome, 314 polysomnography, 314 postoperative, 756–757 protrusor muscle, 330 radial forces, axial forces, interaction between, 323 retrusor muscle, 330 sedation, with midazolam, 335 sensory receptor dysfunction, 329 soft tissue, crowding of, 322 Starling resistor, 318–319 symptoms of, 315–316 therapy, 333–335 upper airway anatomy, alterations in, 316 upper airway collapsibility, 320–322, 325 upper airway dilator muscles, activation of, 327 velopharyngeal muscles, 333 wakefulness, 326 Obstructive sleep apnea-hypopnea syndrome, 399–402 epidemiology, 384, 399 eucapnic obesity, 397 genetics of, 384–385 hypercapnic obesity, 398 nasal mechanical devices, 409–412 with obesity, 399–402 pathogenesis, 401 postoperative, 758 in recovery from anesthesia, 738–739 treatment, 408–412 weight loss, effect of, 408 Olprinone, 755 Ondine’s curse. See Congenital central hypoventilation syndrome Opioids, 103–105, 108, 113, 122, 161, 226, 334, 364, 425–430, 440–490, 514, 516, 524, 526–528, 537–538, 540–557, 577–579, 583, 672, 741–742, 750–754, 758, 761, 784. See also specific opioid with benzodiazepines, 553–554 epidural, 465 intramuscular, 447–448, 753, 760 with ketamine, 556–557 overdose, 450 postoperative, 429 respiratory depression, 442 gender differences in, 444–445
Subject Index Opioids (continued ) postoperative, 445 sleep disturbance, 760 tranquilizers combined with, 555–556 Oropharyngeal muscles, 333 in obstructive sleep apnea, 333 Oscillator model for central anesthetic effects on respiratory rate, 608 Overlap syndrome, in obesity, 412 Oxygenation, 755 Oxyhemoglobin desaturations, 477–478 Ozone, stimulation, 240–241 Pacemaker, 83, 298, 586, 590, 598 Pain, 51, 74, 76–78, 80, 82, 104, 107, 109, 122–123, 243, 247, 425–516, 527, 553, 556, 671–674, 677, 751, 753–754, 760. See also specific analgesic acute, 131, 434, 436, 438, 447–448, 451, 468–469, 473, 489–490, 672, 752 adverse outcomes, link between, 436 cardiovascular function with, 437–438 chronic, 42, 438, 461, 465 dynamic, 439–440 effects of, 438–454 opioids, 440–454 postoperative, 751–752 control of breathing, 751–752 respiratory dysfunction, 426–433 pulmonary function, effects of pain on, 436–437 stress response, effects of pain on, 433–436 surgical trauma, effects of, 433–438 ventilatory effect, 438–454 Palate, general anesthesia, effect on, 692 Panic disorder, breathing patterns with, 77–78 Para-chlorophenylalanine, 192–193 Paradoxic breathing, in recovery from anesthesia, 741 Paralysis, 699–700 Parasternal intercostal, with anesthesia, 701–704 Paravertebral block, 461 hypotension, 461 oxygenation, 461 patient-controlled analgesia, morphine, 461 peak expiratory flow rate, 461 thoracotomy, 461
841 Pathophysiology, 317–322 Patient-controlled analgesia, 107, 383, 447–450, 453–454, 461–462, 473, 482, 486–487, 490 epidural, 436, 462, 490 Peak expiratory flow rate, postoperative, 431 Pentazocine, 464, 473–474 Pentobarbital, 527–528, 579, 584, 586–593, 668, 695–696, 710, 743 hypoxia, 528 intramuscular, 527–528 moderate sedation, 527 oral, 527 Perception of music, effect of, 82–84 Periaqueductal gray matter, opioids and, 442 Perioperative analgesia. See also specific analgesic agent epidural, 484–487 morbidity, 484 pulmonary outcome, 483–487 regional analgesia, 485 respiratory depression, 486 respiratory failure, 486–487 thoracic epidural analgesia, 486 thoracic surgery, 484 upper abdominal surgery, 484 Perioperative myocardial infarction, 757 Peripheral chemoreceptors, 3–19, 601 acetylcholine, 9 carotid body, 3–5 cellular organization, 4–6 glucose sensing in, 12 innervation, 4–6 neuromodulation in, 12–15 chemoreceptor clusters, 4–6 co-culture model, 9 effect of serotonin, 13 fast-acting neurotransmitters, 8–11 chemosensory processing, 8–11 functional genomic approach, 15 gamma-aminobutyric acid, 13 glucose sensing in carotid body, 12 heme proteins, 8 hypoxia, closure by, 8 hypoxic chemotransmission, 11 principal mechanism underlying, 11 large conductance potassium channel, 6–8 blockade of, 7–8 local circuits, role for, 6 mitochondrial involvement, 8 neuromodulation in carotid body, 12–15
842 Peripheral chemoreceptors (continued ) O2-sensitive potassium channels, 6–8 postsynaptic depolarization, recording of, 9–11 presynaptic effects, 12–13 purinergic receptors, molecular identity of, 14–15 regulation of type I cell function, role in, 13–14 sinus nerve, carotid, 3–4 tandem pore acid-sensitive potassium channel (TASK), 6, 8, 13, 46–47, 52, 73–75, 86, 89, 262, 296, 514, 590–591, 627, 664, 673 type I cell function regulation, 13–14 Peristaltic activity, pain and, 438 Pharmacodynamic interaction models, 134–137 mechanism-based approach, 135–136 modeling interaction, 136–137 model selection, 137–140 Richards model, 134–135 Pharyngeal obstruction analgesia with opioids, 741 mechanisms of, 739 positions, effect of, 744 during recovery from anesthesia, 738–745 residual muscle paralysis, 742 surgical, anesthetic interventions, effects of, 742 Pharyngeal receptors, 227 Phasic activity of expiratory muscles, 699 Phasic rapid eye movement, postoperative, 758 Phencyclidine, 591–593 Phenergan, 555 Phenylephrine, 798 Physiogenetic map of ventilatory behavior, 279–282 Physostigmine, 521, 527, 552–553 Pickwickian syndrome, 314, 398 Picoejection, 605, 607, 615–619 Picrotoxin, 606–607, 613–615 Pillow use, pharyngeal obstruction and, 744–745 Plasticity defined, 156 respiratory, modulation, contrasted, 156 Platelet-derived growth factor beta, 162 Pleiotropy, inheritance of ventilatory traits, 280
Subject Index Pleural effusions, postoperative, 428 Pneumonia, postoperative, 428, 435, 754–755 Pneumothorax, postoperative, 428 Poikilocapnia, 118, 169, 178, 358–359, 655, 674, 782 Polysomnography, 314 in obesity, 396 in obstructive sleep apnea, 314 with postoperative hypoemia, 764 Positive end-expiratory pressure, 572, 599–600, 716 Post-hypoxia ventilatory behavior, 162–163, 170, 197 genetic transmission of, 276 Post-inspiratory activity, effect of anesthesia on, 705–706, 709 Post-nasal drip syndrome, 248 Postoperative functional residual capacity, 752–753 Postoperative immobility, effects of, 435 Postoperative pain. See Pain Postoperative pulmonary dysfunction, 428–432 Postoperative respiratory dysfunction, 426–433 factors contributing to, 426 general anesthesia, 427 neuraxial anesthesia, 427 regional anesthesia, 427–428 Postoperative upper airway function anesthesia recovery, 738–749 chemical control of breathing, anesthesia recovery, 750–751 lung function recovery, after anesthesia, 752–754 nocturnal hypoxemia, anesthesia recovery, 755–763 Postsynaptic depolarization, recording of, 9–11 Potassium channel, 30, 47, 365, 481, 589–590, 626–630, 664, 666, 670, 781. See also specific potassium channel Pre-Bo¨tzinger complex, 598 Premotor neuron, 576, 581–583, 598, 601–603, 607–608, 613–615, 617, 620, 622–623, 626, 628, 630 Prenatal hypoxia, respiratory plasticity and, 175–176 Prochlorperazine, 555–556
Subject Index Progestational steroids, 374 Promethazine, 555 Proopiomelanocortin, 264–265, 280 gene, 265 Propofol, 532–536, 576, 584, 587, 589–594, 596, 676, 696, 745, 754 acute hypoxic response, 535–536 airway obstruction, 536 deep sedation, 532 hypoxic ventilatory depression, 535–536 hypoxic ventilatory drive, 535 with ketamine, 557 propofol, intravenous, effect on ventilatory drive, 533–534 pulmonary ventilation, monitoring of, 536 remifentanil, interaction, 148–151 tidal volume, decrease of, 532 Protein kinase C, 161–162 Protrusor muscle, 330 in obstructive sleep apnea, 330 Pseudo-steady-state pharmacodynamics methods, 120 Psychological influences on breathing, 76–79 dimension model, 77–78 dorsal neural network, 79 emotion perception, cerebral correlates of, 78–79 expression of emotion, effect on changes in breathing, 76–77 psychological influences, functional role, 78 self-regulation of emotion, involvement in, 78 specific breathing behaviors, distinguishment of psychological derangements, 77–78 specificity model, 77–78 ventral neural network, 79 Pulmonary afferents, rate changes, 609 Pulmonary aspiration, 244–246 defensive airway reflexes, 246 lung abscess, 245 perioperative periods, complication observed during, 244–245 Pulmonary atelectasis, postoperative, 752 Pulmonary edema, 89, 117–118, 134, 262, 358–360, 376, 520, 528, 531, 544, 603, 622, 688, 690, 692, 698–701, 716–720, 722, 801. See also High-altitude pulmonary edema
843 Pulmonary function, effects of pain on, 436– 437 Pulmonary gas exchange, alteration of, 443 Pulmonary hypertension, 375 Pulmonary stretch receptors, 580, 610–612 Pulse oximetry, 107–108, 558 Purinergic receptors, molecular identity of, 14–15 Purinoceptor agonists, antagonists, 802–804 Quantification in drug pharmacodynamics, 108–109 Quiet breathing anesthetic effects on, 706–707 effect of anesthesia on, 704 Radial forces, axial forces, interaction between, 323 Rapid eye movement postoperative, 758–762 rebound, postoperative, 761–762 suppression, postoperative, 760–761 Rapidly adapting receptors, 236 Read’s rebreathing technique, 49, 110–113, 658–659. See also Rebreathing technique Rebreathing technique, 553–554, 599 advantage of, 115 refinement of, 113 Recovery from anesthesia, 737–777. See also Anesthesia Reflex swallowing, 232 Regional analgesia, 454–473 Regional anesthesia, 425–512 Remifentanil, 117, 148, 150–152, 450, 550– 551 dual-isohypercapnic technique, 550 hypoventilation, hypoxemia resulting from, 551 hypoxia, 550 isohypercapnic technique, 550 propofol, interaction, 148–151 ultra-short-duration opioid, 550 Renin-angiotensin system, 800–801 exercise and, 800 Respiration. See specific aspect of Respiratory neurons, 581–583, 604–605, 613–622 Respiratory neuroplasticity, 155–223. See also Neuroplasticity, respiratory cervical hemisection contralateral effects, 196
844 Respiratory neuroplasticity (continued ) supraspinal effects, 196 crossed phrenic pathways, spontaneous motor recovery, 192–193 developmental, 174–187 hypercapnia, 182–184 hyperoxia, 179 intermittent neonatal hypoxia, 178–179 metabolism, 184–185 neonatal maternal separation, 186–187 prenatal hypoxia, 175–176 sensory denervation, 185–186 sustained neonatal hypoxia, 176–178 hypercapnia, 172–174 acute intermittent hypercapnia, 172 acute sustained hypercapnia, 172 chronic intermittent hypercapnia, 174 chronic sustained hypercapnia, 173 hyperoxia, 171–172 hypoxia, 158–171 acute, 159 brief intermittent, 163 chronic intermittent, 168 hypoxic ventilatory depression, 161–162 with long-duration hypoxia, 170–171 with moderate-duration hypoxia, 170 post-hypoxia frequency decline, 162–169 with short-duration hypoxia, 169 short-term depression, 162–169 short-term potentiation, 159–161 sustained, 167 location of, 156–158 modulation, contrasted, 156 respiratory gases inducing, 158–174 sex hormones, 187 spinal cord injury, 187–198 cervical contusion injuries, 196–197 crossed phrenic pathways, 192–195 dorsal rhizotomy, 197–198 hemisection, 189–192 serotonin, 192 Respiratory neurotransmission, anesthetic effect, 613–629 Respiratory premotor neurons, 613–622 Response surface modeling, drug interactions, 133–153
Subject Index Akaike’s information-theoretic criterion, 137–138, 142–143 bootstrap analysis, 140–143 propofol, remifentanil, 148–151 sevoflurane, alfentanil, 143–148 mechanism-based approach, 135–136 modeling interaction, 136–137 model selection, 137–140 Monte Carlo simulation, 139 multimodel inference, 137–140 pharmacodynamic interaction models, 134–137 Richards model, 134–135 Resting measurements, in drug pharmacodynamics, 109–110 Retrusor muscle, 330 in obstructive sleep apnea, 330 Rhizotomy, 166, 188, 197–198 Rhythm generation in adult mammal, functional model for, 601 Rib cage effect of anesthesia on, 688–689 expansion of, effect of anesthesia, 690 Richards model, pharmacodynamic interaction, 134–135 Rostral ventral medulla lateral medulla, chemosensitive area, 53–55 opioids and, 442 Scalene, 702–703 Schizophrenia, breathing patterns with, 77 Sedalgesia, 513 Sedation levels, 335 deep, 514–515 minimal, 514–515 moderate, 514–515 Sedatives, 516–540. See also specific sedative minimizing respiratory risks of, 557–559 dose titration, 557–558 supplemental oxygen, 558–559 in mountain sickness, 373–374 ventilatory effect of, 516–540 Self-regulation of emotion, involvement in, 78 Sensory denervation, respiratory plasticity and, 185–186 Serotonin, 4, 13, 43–45, 157, 161, 163–168, 170, 186, 190–195, 197–198, 226, 236–237, 277, 362, 367, 376, 473, 475, 572, 607–608, 626–629, 673–674
Subject Index Serotonin (continued ) receptor, 166, 170, 186, 192, 195, 197 Serum cholesterol levels, in obesity, 400 Sevoflurane, 133, 143–152, 574–576, 580– 581, 584, 592, 594–595, 608, 617–624, 628, 655, 659–663, 666, 668–669, 671–672, 675–676, 749, 752 alfentanil, interaction, 143–148 Sex hormones, 165, 187, 262 Sexual dimorphism, inheritance of ventilatory traits, 282 Short-duration hypoxia, chronic intermittent hypoxia with, 169 Short-term potentiation of breathing, 674–676 Sleep, 445–446. See also Sleep apnea architecture, postoperative changes in, 758–762 fragmentation, postoperative, 758 hypoxemia during, postoperative, mechanisms of, 762–763 lower arterial oxygen saturation during, 375 periodic breathing during, 374 surgical stress and, 761 Sleep apnea, 248–249, 313–356, 741, 797. See also Obstructive sleep apnea acromegaly, 316 airflow, maximal inspiratory, 321–322 anatomic factors, 322–325 apnea-hypopnea index, 320–321 blood pressure surges, 317 cardiovascular disease, 316–317 central nervous system depressants, 334–335 cervical strap, 332 cervical structures, 322–321 clinical risk factors, 314–317 continuous positive airway pressure, 333, 335 critical pressure, 319–328 lowering of, 333 EMG activity of muscles, 328–330 endocrinopathy, 316 endotracheal extubation in, 743 epidemiology, 314–317 factors, modeling of, 318–320 familial factors in, 263–266, 315 gender and, 315 gene regions in human associated with, 264
845 genioglossus decreases in activity, 329–330 role of, 330–332 with hypercapnia, obesity and, 402–407 hypothyroidism, 316 monitoring technology, 314 neuromuscular compensatory mechanisms, 325–327, 333–334 neuromuscular factors, 325–333 neuromuscular reflexes, 325 upper airway muscles, 330 with obesity, 313–316, 323–325, 399–407 oropharyngeal muscles, 333 pathogenesis, 317–333 pathophysiology, 317–322 peri-operative setting, 334–335 pharynx, site of obstruction, 317–319 Pickwickian syndrome, 314 polysomnography, 314 postoperative, 756–757 protrusor muscle, 330 radial forces, axial forces, interaction between, 323 retrusor muscle, 330 sedation, with midazolam, 335 sensory receptor dysfunction, 329 soft tissue, crowding of, 322 Starling resistor, 318–319 symptoms of, 315–316 therapy, 333–335 upper airway anatomy, alterations in, 316 upper airway collapsibility, 320–322, 325 upper airway dilator muscles, activation of, 327 velopharyngeal muscles, 333 wakefulness, 326 Sleep apnea-hypopnea syndrome, 399 epidemiology, 384, 399 eucapnic obesity, 397 genetics of, 384–385 hypercapnic obesity, 398 nasal mechanical devices, 409–412 with obesity, 399–402 pathogenesis, 401 postoperative, 758 in recovery from anesthesia, 738–739 treatment, 408–412 weight loss, effect of, 408 Slowly adapting receptors, 235–236, 238–240 Slow-wave sleep, 84–85 Small conductance potassium channel, 590 Smooth muscle, reflex relaxation of, 238
846 Sniffing position, pharyngeal obstruction and, 744 S-nitrosothiol, 676 Speech, tonic excitatory drive, 87–88 Spinal cord injury, respiratory plasticity and, 187–198 cervical contusion injuries, 196–197 cervical hemisection contralateral effects, 196 supraspinal effects, 196 crossed phrenic motor output, strengthening, 193–195 crossed phrenic pathways, 192 spontaneous motor recovery, 192–193 dorsal rhizotomy, 197–198 hemisection, 189–192 serotonin, 192 Spinal hemisection, respiratory plasticity and, 189–192 breathing at rest, 189–190 crossed phrenic phenomenon, 190–192 minute ventilation, 189 Splinting, postoperative, 435 Sprague-Dawley rat, inheritance of ventilatory traits, 271–273, 278, 280 Starling resistor, 318–319 Stress response, effects of pain on, 433–436 Stridor, after extubation, 745 Strychnine, 601, 607, 613 Substance P, 4, 160, 223, 229, 243, 362, 473, 626–627, 629, 781 Succinate dehydrogenase, 666 Sudden infant death syndrome, 158, 233, 296, 757 Sufentanil, 447, 451, 456, 466–467, 469, 471– 473, 480, 802 Supplemental oxygen, 558–559 Suprapontine control of breathing, 71–102 behavioral control, automatic control, interaction between, 86–88 inhibitory drives, 84–86 involuntary emotional influences, 75–84 learned respiratory behaviors, 88–90 music perception, 82–84 psychological influences, 76–79 respiratory discomfort perception, 79–82 tonic excitatory drives, 84–86 volitional control, 72–75 Surgical stress, postoperative sleep disturbance after, 761 Surgical trauma effects of, 433–438
Subject Index postoperative, 429–432 Sustained hypoxia responses, 363–369 Sustained neonatal hypoxia, respiratory plasticity and, 176–178 Swallowing states, 232 Synaptosome, 584–585, 588–589 Tachycardia, 435, 437 Tachypnea, 431, 608–609 Tandem pore acid-sensitive potassium channel (TASK), 6, 8, 13, 46–47, 52, 73–75, 86, 89, 262, 296, 514, 590–591, 627–628, 664, 673–674 Tandem pore acid-sensitive potassium channel-1 (TASK-1), 41, 44–45, 628 Tandem pore weak inward rectifying potassium channel-1 (TWIK-1), 6, 591, 664 TASK channel. See Tandem pore acidsensitive potassium channel (TASK) TEIK-1-related potassium channel-1 (TREK-1), 591, 664 Temazepam, in mountain sickness, 373–374 Tetrodotoxin, 38, 572, 587–588, 599, 607, 628 Theophylline, 162, 193–194, 373, 380, 803–804 Thiopental, 529–532, 535–536, 579, 584, 586–590, 625–626, 696–697, 703–704, 710–711, 713, 721, 754 airway musculature, effect on, 531 deep sedation, 529 general anesthesia, 529 hypoxia, 531 hypoxic ventilatory response, effect on, 531 intravenous, effect on ventilatory drive, 530 methohexital, 531 rebreathing technique, 529 Thoracic curvature, functional residual capacity with, 698 Thorax, effect of anesthesia on, 688–689 Thorazine, 555 Thromboembolic phenomena, 437–438 Thyrotropin-releasing hormone, 44–45, 608, 626–627, 629 Tidal volume effect of anesthesia on, 709 effect of pain, 436
Subject Index Time course, drug effect, 116–117 Tonic diaphragm activity, cephalad diaphragm displacement, anesthesia effects on, 699 Tonic excitatory drive, 84–86, 601–602 reduction of, 608 Tonic inhibitory drives, 84–86 Tonic respiratory drive, 599 TRAAK channel, 591, 664 Trachea, irritation of, 240 Tracheal carina, stimulation of, 240 Tracheal intubation continuous positive airway pressure, 747–748 coughing during, 747 cuff-leak test, 747 duration of, 747 trauma at, 747 Tracheal receptors, 227 Tracheal tube, tight-fitting, 747 Tramadol, 474–475 Tranquilizers, 513 with opioids, 555–556 Transversus abdominis, effect of anesthesia on, 701–703 TREK-1 channel, 591, 664 Trichloroethylene, 611 TWIK-1 channel, 591, 664 Tyrosine hydroxylase, 4–6, 366 Upper airway anatomic alterations, in obstructive sleep apnea, 316 collapsibility, 320–322, 325 dilator muscles, activation of, 327 function, recovery from anesthesia, 738–749 mechanics, effect of anesthesia, 712–714 reflex responses of, 229–234 airway, defense, protection, 229–233 apnea, 231 larynx, 230–231 maintenance of airway patency, 233–234 nose, 229 pharynx, 232 Upper airway obstruction, 313–356 acromegaly, 316 airflow, maximal inspiratory, 321–322 anatomic factors, 322–325 apnea-hypopnea index, 320–321 blood pressure surges, 317
847 cardiovascular disease, 316–317 central nervous system depressants, 334–335 cervical strap, 332 cervical structures, 322–321 clinical risk factors, 314–317 collapsibility, 763 continuous positive airway pressure, 333, 335 critical pressure, 319–328 lowering of, 333 EMG activity of muscles, 328–330 endocrinopathy, 316 epidemiology, 314–317 factors, modeling of, 318–320 familial aggregation, 315 gender and, 315 genioglossus decrease in activity, 329–330 role of, 330–332 hypothyroidism, 316 monitoring technology, 314 neuromuscular factors, 325–333 neuromuscular reflexes, 325 upper airway muscles, 330 neuromuscular mechanisms, compensatory increases, 325–327, 333–334 obesity, 313–316, 323–325 oropharyngeal muscles, 333 pathogenesis, 317–333 pathophysiology, 317–322 peri-operative setting, 334–335 pharynx, site of obstruction, 317–319 Pickwickian syndrome, 314 polysomnography, 314 protrusor muscle, 330 radial forces, axial forces, interaction between, 323 retrusor muscle, 330 sedation, with midazolam, 335 sensory receptor dysfunction, 329 soft tissue, crowding of, 322 Starling resistor, 318–319 symptoms of, 315–316 therapy, 333–335 velopharyngeal muscles, 333 wakefulness, 326 Urinary retention, pain and, 438 Vagotomy, 580 Vascular endothelial growth factor, 181
848 Vecuronium, 122, 742–743, 750, 782–785, 787 Venous thrombosis, postoperative, 435 Ventilatory acclimatization, 167, 170, 358–361, 363, 365, 367, 370, 372–373 to hypoxia, 167–168, 175, 199, 359–361, 365–368, 383–384, 386, 388, 390, 392, 395–396, 398–400, 402, 404–408, 410, 412, 804, 806 Ventilatory control disorders with obesity, 399–407 Ventilatory deacclimatization, 367–368 Ventilatory pharmacological effect measurement, 103–131 airway gas concentration of drug, 106–108 airway pressure alterations, 120–122 flow limitation, 122 genioglossal EMG, 122 negative airway pressure, 122 resistance, 122 control of breathing, 109–110 dual-isohypercapnic technique, 117 dynamic end-tidal forcing-type system test, 118 dynamic hypercapnic response model, 120 exercise, 122 hyperbola model, 118 hypercapnic ventilatory response, 110–117 hypoxic ventilatory depression, 120 hypoxic ventilatory response, 117–120 inductance plethysmography, 108 interpretation, complexity of, 109–110 isocapnic conditions, maintenance of, 117–118 linear-in-saturation model, 118 modeling technique, 117 pain, 122–123 plasma concentration response curve, 108 pseudo-steady-state methods, 120 pulse oximetry, 107–108 quantification, drug pharmacodynamics, 108–109 Read’s rebreathing technique, 110–113 rebreathing technique advantage of, 115 refinement of, 113 resting measurements, 109–110
Subject Index steady-state technique, hypercapnic ventilatory response, 111–113 time course, drug effect, 116–117 Ventilatory response to CO2, 47–50 dynamic end-tidal forcing, 50 hyperoxia to separate peripheral, central inputs, 47 recovery of CO2 response after, 49 ventilatory/phrenic nerve response to CO2, abolition of, 48 ventral medullary surface areas, cooling of, 48 ventriculocisternal perfusion, 47 Ventilatory trait inheritance, 261–266 gene effects in rodent models, 276–279 physiogenetic map, ventilatory behavior, 279–282 in rodents, 271–273 in small animals, 266–271 strength of inheritance, 274–276 Ventral medullary raphe region, as chemosensitive area, 55 Ventral neural network, 79 Ventral respiratory group, 28, 37, 157, 194, 443–444, 581, 583, 597–598, 601, 613–615, 623, 651 Ventriculocisternal perfusion, 47 Verapamil, 802 Vital capacity, 111, 113, 116, 303, 388–392, 398, 406–407, 411–412, 428–429, 431–436, 457, 460, 463–464, 466, 487, 790 Volatile anesthetics carotid body mediated ventilatory response, 658 effects of, 574, 615 minimum alveolar concentration of, 655 at subanesthetic concentrations, 658 Volitional control of breathing, 72–75 activation sites, instruments to detect, 75 awareness of breathing, 73 distributed neural network, 75 locked-in syndrome, 74 maximum voluntary ventilation, 73 motor cortex, neural origin, 74–75 overriding metabolic demands, advantage of, 72–73 voluntary drives to breathe, role of, 74 Voltage-gated calcium channel, 584, 586, 588 Voltage-gated cation channel, 596 Voltage-gated ion channel, 584–591
Subject Index Voltage-gated potassium channel, 589–590 Voltage-gated sodium channel, 588 Waist-hip ratio, in obesity, 400 Wakefulness drive, 52, 76, 83–85, 105, 109–110 Weight loss effect on ventilatory response, 408–409 improvement in lung mechanics with, 388–389
849 Wistar rats, inheritance of ventilatory traits, 272 Xenon, 392, 592–593 Zolpidem, with acute mountain sickness, 373–374 Zucker rats, heritability estimates, 276
